Photosynthesis Photobiochemistry and Photobiophysics (Advances in Photosynthesis and Respiration)

Photosynthesis Photobiochemistry and Photobiophysics Advances in Photosynthesis VOLUME 10 Series Editor: GOVINDJEE U...

Author: Bacon Ke

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Photosynthesis Photobiochemistry and Photobiophysics

Advances in Photosynthesis VOLUME 10

Series Editor: GOVINDJEE University of Illinois, Urbana, Illinois, U.S.A.

Consulting Editors: Jan AMESZ, Leiden, The Netherlands Eva-Mari ARO, Turku, Finland James BARBER, London, United Kingdom Robert E. BLANKENSHIP, Tempe, Arizona, U.S.A. Norio MURATA, Okazaki, Japan Donald R. ORT, Urbana, Illinois, U.S.A.

Advances in Photosynthesis is an ambitious book series seeking to provide a comprehensive and state-of-the-art account of photosynthesis research. Photosynthesis is the process by which higher plants, algae and certain species of bacteria transform and store solar energy in the form of energy-rich organic molecules. These compounds are in turn used as the energy source for all growth and reproduction in these organisms. As such, virtually all life on the planet ultimately depends on photosynthetic energy conversion. This series of books spans topics from physics to agronomy, from femtosecond reactions to season long production, from the photophysics of reaction centers to the physiology of whole organisms, and from X-ray crystallography of proteins to the morphology of intact plants. The intent of this series of publications is to offer beginning researchers, advanced undergraduate students, graduate students, and even research specialists a comprehensive current picture of the remarkable advances across the full scope of photosynthesis research.

The titles published in this series are listed at the end of this volume and those of forthcoming volumes on the back cover.

Photosynthesis Photobiochemistry and Photobiophysics

Bacon Ke Walnut Creek, CA, U.S.A.

KLUWER ACADEMIC PUBLISHERS NEW YORK, BOSTON, DORDRECHT, LONDON, MOSCOW

eBook ISBN: Print ISBN:

0-306-48136-7 0-7923-6334-5

©2003 Kluwer Academic Publishers New York, Boston, Dordrecht, London, Moscow Print ©2001 Kluwer Academic Publishers Dordrecht All rights reserved No part of this eBook may be reproduced or transmitted in any form or by any means, electronic, mechanical, recording, or otherwise, without written consent from the Publisher Created in the United States of America Visit Kluwer Online at: and Kluwer's eBookstore at:

http://kluweronline.com http://ebooks.kluweronline.com

To Keiko

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Editorial: Advances in Photosynthesis and Respiration I grew up in the world of photosynthesis. My studies in the field began in 1953, when I was a Master’s of Science student in Shri Ranjan’s class at Allahabad University, India. Ranjan had not only worked in the laboratory of F. F. Blackman, which is known for the law of limiting reactions in photosynthesis, but had also done research in respiration. At that time, I read Volume I (published in 1945) and the first part of Volume II (published in 1951) of Eugene Rabinowitch’s masterpiece, Photosynthesis and Related Processes. It was only after 1957, when I came to the University of Illinois at Urbana-Champaign to work with Robert Emerson, that I read the rest of the masterpiece – Volume II, part 2 (1956). This book (2088 pages long) made a tremendous impact on the research of many. Today, it would be impossible for one author to write a comparable, in-depth book covering all aspects of photosynthesis –the subject has grown too complex. Between 1990 and 2000, according to Current Contents, 11,576 papers were published on photosynthesis. Of those, 3,796 were on photosystem II; 1,059 on photosystem I, 2,044 on chlorophyll a fluorescence, 1,270 on ATP synthase; 1,435 on oxygen evolution, and 1,050 on RIBISCO. Further, a search on the Internet through turned up 106,000 on photosynthesis, 8,190 hits on photosystem II, 8,290 hits on chlorophyll fluorescence, 17,400 hits on ATP synthase, and 5,550 on RUBISCO. Wow! In the early 1990s, I initiated the series “Advances in Photosynthesis” with Kluwer Academic Publishers. My earlier experience in writing and editing books (“Photosynthesis” with E. Rabinowitch, John Wiley & Sons, 1969; “Bioenergetics of Photosynthesis,” 1975; “Photosynthesis,” Vols. I and II, 1982; and “Light Emission by Plants and Bacteria,” co-edited with Jan Amesz and Dave Fork, 1986; Academic Press) was of great help in this new endeavor. An earlier series entitled “Topics in Photosynthesis,” edited by J. Barber (Elsevier, the Netherlands), 1976 to early 1990s, had already pioneered the practice of multi-authored books in photosynthesis. Although “Advances in Photosynthesis” was launched before this earlier series had ended, both series proved valuable in this rapidly expanding field. With the help of Larry Orr at Arizona State University, Tempe, and the assistance of Ad Plaizier, Gilles Jonker, Natasha Bonnevalle, Jacco Flipsen, and Gloria Verhey at Kluwer Academic Publishers, we have published the following nine books in the “Advances in Photosynthesis” series. (1) Molecular Biology of Cyanobacteria (D. A. Bryant, editor, 1994); (2) Anoxygenic Photosynthetic Bacteria (R. E. Blankenship, M. T. Madigan and C. E. Bauer, editors, 1995); (3) Biophysical Techniques in Photosynthesis (J. Amesz and A. J. Hoff, editors, 1996); (4) Oxygenic Photosynthesis: The Light Reactions (D. R. Ort and C. F. Yocum, editors, 1996); (5) Photosynthesis and the Environment (N. R. Baker, editor, 1996); (6) Lipids in Photosynthesis: Structure, Function and Genetics (P.-A. Siegenthaler and N. Murata, editors, 1998); (7) The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas (J-D. Rochaix, M. Goldschmidt-Clermont and Sabeeha Merchant, editors, 1998); (8) The Photochemistry of Carotenoids (H. A. Frank, A. J. Young, G. Britton and R. J. Cogdell, editors, 1999); and (9) Photosynthesis: Physiology and Metabolism (R. C. Leegood, T. D. Sharkey and Susanne von Caemmerer, editors, 2000). vii

See for further information and to order these books. Please note that members of the International Society of Photosynthesis Research, ISPR () receive special discounts. Although I continue to believe in the value of multi-authored books in this series, I decided that this precedent could be broken when I discovered that Bacon Ke had written a marvelous manuscript on photosynthesis in the tradition of Eugene Rabinowitch. I have read Ke’s work, “Photosynthesis: Photobiochemistry and Photobiophysics,” and believe it to be an important addition to the “Advances in Photosynthesis” series. Through this book I have learned to appreciate how the kinetics of various reactions, measured by difference absorption spectroscopy and by electron spin resonance, have led to the current understanding of the pathway of electron transport in both plant and bacterial photosynthesis. Bacon has woven into the book the atomic structure of the various protein complexes with their function, in both a scientific and artistic manner, making it unique in dealing with the biochemistry and biophysics of plant, cyanobacterial, algal, and bacterial photosynthesis. In telling the story of photosynthesis, Bacon has dealt with the time sequence of reactions from picoseconds to milliseconds in a beautiful manner. Using the scale = -log(time in seconds), from Martin Kamen’s 1963 book), Bacon has mostly covered the decades from 12 (picosecond) to 3 (millisecond). This book will be considered a classic in its field and will serve all beginning students and researchers in the area of the light reactions of photosynthesis, up to the production of NADH or NADPH and ATP. It is the first book in the “Advances in Photosynthesis” series that can be used as a text book in a graduate or upper division undergraduate course on photosynthesis; we will only need to supplement it with a book on the dark reactions, such as volume 9 of the series, to cover the entire range of photosynthesis. This great book by Bacon Ke is volume 10 in the “Advances in Photosynthesis” series. The format of this single-authored book is intentionally different from the previous nine multi-authored books. We hope to continue using both formats in the future. In Volume 11, we will make several important changes to the series. The topic of “Respiration” will be added, and, thus, the title of the series will be changed to “Advances in Photosynthesis and Respiration.” We will also have a new board of consulting editors. Outgoing editors Jan Amesz, The Netherlands; Eva-Mari Aro, Finland; James Barber, UK; Robert E. Blankenship, USA; Norio Murata, Japan; and Donald R. Ort, USA, will be replaced in volume 11 by Eva-Mari Aro, Finland; Christine Foyer, UK; Elisabeth Gantt, USA; John H. Golbeck, USA; Susan Golden, USA; Wolfgang Junge, Germany; Hartmut Michel, Germany; and Kimiyuki Satoh, Japan. Eva-Mari Aro (Finland), who joined us only in Vol. 3, has graciously agreed to serve till Vol. 12, when a new member in the area of respiration will be appointed. William Ogren served on the board of consulting editors for volume 1. I am particularly thankful to Jan Amesz, Eva-Mari Aro, Bob Blankenship, Norio Murata, and Don Ort for taking an active role in editing the various volumes in the series. Further, the following new members have already agreed to edit future volumes: Christine Foyer, John Golbeck, Wolfgang Junge, and Kimiyuki Satoh. I thank all the outgoing consulting board members, the past editors and the authors, and welcome all the incoming board members, the editors and the authors of the future volumes. I would also like to thank Prasanna Mohanty for making many suggestions for both the current and future book series. Prasanna has also written detailed reviews of our books in several Indian journals, bringing information to the photosynthesis community in the developing world. It is my dream that some day we will increase the access to these books by producing a version that all graduate students throughout the World can afford to purchase. viii

The future scope of our series will reflect the recognition that photosynthesis and respiration are intertwined with respect to both the protein complexes involved and to the entire bioenergetic machinery of all life. “Advances in Photosynthesis and Respiration” will be a book series providing a comprehensive and state-of-the-art account of research in photosynthesis and respiration. Photosynthesis is the process by which higher plants, algae, and certain species of bacteria transform and store solar energy in the form of energy-rich organic molecules. These compounds are in turn used as the energy source for all growth and reproduction in these and almost all other organisms. As such, virtually all life on the planet ultimately depends on photosynthetic energy conversion. Respiration, which occurs in mitochondrial and bacterial membranes, utilizes energy present in organic molecules to fuel a wide range of metabolic reactions critical for cell growth and development. In addition, many photosynthetic organisms engage in energetically wasteful photorespiration that begins in the chloroplast with an oxygenation reaction catalyzed by the same enzyme responsible for capturing carbon dioxide in photosynthesis. This series of books spans topics from physics to agronomy and medicine, from femtosecond processes to season long production, from the photophysics of reaction centers, through the electrochemistry of intermediate electron transfer, to the physiology of whole organisms, and from X-ray crystallography of proteins to the morphology of organelles and intact organisms. The intent of the series is to offer beginning researchers, graduate students, advanced undergraduate students, and even research specialists, a comprehensive, up-to-date picture of the remarkable advances across the full scope of research on bioenergetics and carbon metabolism. The readers of the current series are encouraged to watch for the forthcoming books: Regulatory Aspects of Photosynthesis (volume 11; Editors: E.-M. Aro and B. Andersson); Light-harvesting Antennas in Photosynthesis (Editors: B. R. Green and W. W. Parson); Photosynthesis in Algae (Editors: A. W. D. Larkum, S. Douglas, and J. A. Raven); Photosynthetic Nitrogen Assimilation and Associated Organic Acid Metabolism (Editors: C. H. Foyer and G. Noctor); (5) Clorophyll Fluorescence: A Signature of Photosynthesis (Editors: G. Papageorgiou and Govindjee); and, (6) Photosystem II: The Water/Plastoquinone Oxido-reductase in Photosynthesis (Editors: T. Wydrzynski and K. Satoh).

(1) (2) (3) (4)

In addition to these contracted books, on-going discussions are taking place for books on Photosystem I; Functional Genomics; Protonation and ATP Synthesis; Plant Respiration; The Chlorophylls; Photoinhibition; Global Aspects and Geochemistry; History of Photosynthesis; The Chloroplasts; C-3 and C-4 Plants; and Plant Photosynthetic Production: A Manual of Methods. Both Kluwer ([email protected]) and I ([email protected]) welcome suggestions for titles of new books (especially in the area of respiration), for editors, for authors, and for improvement of this book series in general. In view of the interdisciplinary character of research in photosynthesis and respiration, it is my earnest hope that this series of books will be used in educating students and researchers not only in Plant Sciences, Molecular and Cell Biology, Integrative Biology, Biotechnology, Agricultural Sciences, Microbiology, Biochemistry, and Biophysics, but also in Bioengineering, Chemistry, and Physics. During the many years that I have edited this series, I have often thought of a quote by William Safire, a columnist for the New York Times. He wrote, “To communicate, put your thoughts in order, give them a purpose, use them to persuade, to instruct, to discover, to seduce.” Regardless of the political ix

views of Safire, I like this quotation. I have recognized it as an important goal for writing (and editing). It is in part because of that goal that I thank all the authors of all the books in this series –both for their valuable writing, and for their patience when I made corrections, sometimes to their proofs! Although I will continue to be actively involved in the new series, I would like to take this opportunity to thank my friend John Whitmarsh for his constant support during difficult times of my life, and to thank John, along with Colin Wraight, Don Ort, and Tony Crofts, for providing a unique intellectual environment for my research in photosynthesis at Urbana. Others at the University of Illinois who have collaborated with my laboratory are the late Herbert Gutowsky, Enrico Gratton, and Robert Clegg. It is, however, my graduate students and post-doctorals who did all the research and they are the ones who were responsible for my decision to undertake the cause of education through the AIPH series. I am highly indebted to Rajni Govindjee, my constant companion since 1957, for her love, support and tolerance, especially of the sometimes intrusive nature of my work (such as being engrossed in e-mail when dinner is ready). I also feel a deep sense ofgratitude to our children Sanjay Govindjee and Anita GovindjeeChristiansen, to their spouses Marilyn Govindjee and Morten Christiansen, and to our grandchildren Sunita Christiansen, Arjun Govindjee, and Rajiv Govindjee.

August 15, 2000 Govindjee Series Editor, Advances in Photosynthesis and Respiration University of Illinois at Urbana-Champaign 265 Morrill Hall, 505 South Goodwin Avenue Urbana, IL 61801-3707, U.S.A. Fax: 217-244-7246 E-mail: [email protected] URL: http://www.life.uiuc.edu/govindjee

x

Contents at a Glance Overview 1.

Photosynthesis: An Overview

1

Bacterial Photosynthesis 2. 3. 4. 5. 6. 7. 8. 9. 10.

The Bacterial Photosynthetic Reaction Center: Chemical Composition and Crystal Structure Light-Harvesting Pigment-Protein Complexes of Photosynthetic Bacteria The Primary Electron Donor (P) of Photosynthetic Bacteria The Stable Primary Electron Acceptor of Photosynthetic Bacteria The Secondary Electron Acceptor of Photosynthetic Bacteria The Early Electron Acceptors of Photosynthetic Bacteria Bacteriochlorophyll and Bacteriopheophytin The Green Bacteria. I. The Light-Harvesting Complex, the Chlorosomes The Green Bacteria. II. The Reaction Center Photochemistry and Electron Transport The Secondary Electron Donor of Photosynthetic Bacteria the Cytochromes

47 65 87 101 111 129 147 159 179

Photosystem II 11. 12. 13. 14. 15. 16. 17.

Photosystem II Introduction The Light-Harvesting Chlorophyll-Protein Complexes of Photosystem II Role of Carotenoids in Photosynthesis Phycobiliproteins and Phycobilisomes The Primary Electron Donor of Photosystem II, P680, and Photoinhibition The Stable Primary Electron Acceptor and the Secondary Electron Acceptor The Transient Intermediate Electron Acceptor of Photosystem II, Pheophytin

199 215 229 251 271 289 305

Oxygen Evolution 18. 19. 20. 21. 22. 23. 24.

Oxygen Evolution Introduction Oxygen Evolution The Role of Manganese Oxygen Evolution UV Absorbance Changes associated with S-state Transitions Oxygen Evolution Extrinsic Polypeptides and Inorganic Ionic Cofactors The Electron Donor to EPR Spectroscopy The Electron Donor to Optical Spectroscopy Charge Recombination in Photosystem II and Thermoluminescence

323 337 355 365 377 397 407

Photosystem I 25. 26. 27. 28. 29. 30. 31. 32. 33. 34.

Photosystem I Introduction 419 Photosystem-l Membrane, Complexes and Crystals 431 Light-Harvesting Chlorophyll-Protein Complexes of Photosystem I 445 The Primary Electron Donor of Photosystem I - P700 463 The Membrane-Bound Iron-Sulfur Proteins (FeS-A and FeS-B): Secondary Electron Acceptors of 479 Photosystem I P430: The Spectral Species Representing the Terminal Electron Acceptor of Photosystem I 505 The Iron-Sulfur Center FeS-X of Photosystem I, the Photosystem-l Core Complex, and Interaction of the FeS-X Domain with FeS-A/FeS-B 527 The Primary Electron Acceptor of Photosystem I 555 The Intermediate Electron Acceptor of Photosystem I – Phylloquinone (Vitamin 579 Mobile Electron Carriers Plastocyanin and Ferredoxin, and Ferredoxin• Reductase 605

Proton Transport and Photophosphorylation Complex and the Related 35. The Interphotosystem 36. Proton Translocation and ATP Synthesis

xi

Complex

635 665


Contents Editorial: Advances in Photosynthesis Series Contents Preface Color Plates

vii

xi xix xxi

Overview 1. Photosynthesis: An Overview

1-46

I. Solar Energy and Photosynthesis II. Absorption of Light Energy and the Fate of the Electronically Excited States III. Light-Harvesting Pigment Molecules IV. Development of Some Modern Concepts of Photosynthesis V. The Thylakoid Membrane of Chloroplasts VI. The Emerson Enhancement Effect and Evidence for Two Photosystems VII. Freeze-Fracture Electron Microscopy of Thylakoid Membranes VIII. Composition of Thylakoid Membranes Lipids, Proteins and Electron Carriers IX. Lateral and Transverse Asymmetry of Thylakoid Membranes X. Evolution of Photosynthetic Reaction Centers

1 4 6 13 19 22 26 30 38 41

Bacterial Photosynthesis 2. The Bacterial Photosynthetic Reaction Center: Chemical Composition and Crystal Structure I. Structure of the Reaction Centers of Photosynthetic Bacteria II. Three-Dimensional Structure of Bacterial Photosynthetic Reaction Centers III. Summary Reaction-Center Structure and Photochemical Reactions

47-64 48 54 62

3. Light-Harvesting Pigment-Protein Complexes of Photosynthetic Bacteria 65-85 I. II. III. IV.

The Light-Harvesting Pigment-Protein Complexes An Early Model for the Light-Harvesting Pigment-Protein Complex Crystal Structure of the Light-Harvesting Complexes of Purple Photosynthetic Bacteria Structure and Function of the Bacterial Photosynthetic Apparatus

4. The Primary Electron Donor (P) of Photosynthetic Bacteria I. II.

Spectral Properties Other Physical Properties The Oxidation-Reduction Potential and the Differential Molar Extinction Coefficient III. Bacteriochlorophyll Epimer being a Constituent of the Primary Electron Donor of Heliobacteria

5. The Stable

Primary ElectronAcceptor

66 69 71 81

87-99 90 96 97

of Photosynthetic Bacteria 101-110

I. Early Spectroscopic Studies of the Stable PrimaryElectron Acceptor II. Functional Role of Ubiquinone and Iron in Photosynthetic Bacteria III. The Oxidation-Reduction Potential of the Stable Primary Electron Acceptor

xiii

102 105 107

6. The Secondary Electron Acceptor I. II. III. IV. V.

of Photosynthetic Bacteria

111-128 112 113 114 117 127

Structure of the Binding Domain and Functional Properties of and Evidence for a Secondary Quinone Electron Acceptor Electron Transfer between the Two Quinones Binary Oscillations Protonation of the Secondary Electron Acceptor Transmembrane Proton Pump and ATP Synthesis A Preliminary Overview

7. The Early Electron Acceptors of Photosynthetic Bacteria Bacteriochlorophyll and Bacteriopheophytin

129-146

in Photochemical Charge Separation I. Discovery of the Involvement of or II. Photochemical Accumulation of the Reduced III. Involvement of the Monomeric BChl in Photochemical Charge Separation

130 133 137

8. The Green Bacteria. I. The Light-Harvesting Complex, the Chlorosomes 147-158 I. II. III. IV. V.

Models for the Reaction Center-Antenna Complex of Green Bacteria Organization of Bacteriochlorophyll-c Molecules in the Rod Elements of Chlorosomes Organization of Chlorophyll Molecules in the Reaction Center-Antenna Complex Pathway and Kinetics of Excitation Energy Transfer in Chlorosomes The BChl a-Protein Complex (the FMO Protein ) in the Reaction Center-Antenna Complex

9. The Green Bacteria. II. The Reaction Center Photochemistry and Electron Transport I. II. III. IV.

159-178

The Green Sulfur Bacteria The Green Filamentous Bacteria The Newly Discovered Photosynthetic Bacteria the Heliobacteria Summary

10. The Secondary Electron Donor of Photosynthetic Bacteria the Cytochromes I. II. III. IV.

149 150 152 153 155

160 168 173 174

179-198

Reduction of Photooxidized Primary Electron Donor by Cytochromes Cytochrome Electron Transfers in the Photosynthetic Bacteria can be Temperature Insensitive Tetraheme Arrangement in the Reaction-Center Complex of Rhodopseudomonas viridis A Reaction Center-Cytochrome c Model System

182 184 186 194

Photosystem II 11. Photosystem II

Introduction

199-214

I. Photosystem II: Protein Components and their Organization II. Structure of the Photosystem-ll Reaction-Center Complex III. Topics on Photosystem II and Oxygen Evolution

200 202 211

12. The Light-Harvesting Chlorophyll-Protein Complexes of Photosystem II 215-228 I. Photosystem-ll Light-Harvesting Chlorophyll-Protein Complexes II. Crystal Structure of LHC II III. Regulatory Roles of LHC II in Thylakoids

xiv

216 219 224

13. Role of Carotenoids in Photosynthesis

229-250

The Light-Harvesting Role of Carotenoids Chlorophyll-Carotenoid-Protein Supramolecular Assemblies Xanthosomes Singlet-Singlet Energy Transfer Dynamics involving Carotenoids in Light-Harvesting The Photoprotective Role of Carotenoids Triplet-Triplet Energy Transfer

I. II. III. IV.

14. Phycobiliproteins and Phycobilisomes

231 233 240 245

251-269

I. Phycobiliproteins II. Phycobilisomes

253 260

15. The Primary Electron Donor of Photosystem II, P680, and Photoinhibition 271-288 I. The Primary Electron Donor of Photosystem II, P680 II Photoinhibition in Photosystem II Consequence of the Strong Oxidizing Power of III. Possible Protective Function of Cytochrome b559 against Photoinhibition

16. The Stable Primary Electron Acceptor Acceptor I. II. III. IV. V. VI. VII.

272 277 282

and the Secondary Electron 289-304

The Stable Primary Electron Acceptor of Photosystem II Absorption-Spectrum Change accompanying the Reduction of Redox Properties of determined by Fluorescence Titration The Two-Electron Gate The Plastoquinone-Nonheme Iron Complex The Binding Sites of and Fe The Herbicide-Binding Protein D1

290 291 294 295 297 299 300

17. The Transient Intermediate Electron Acceptor of Photosystem II, Pheophytin 305-322 Photoaccumulation of Reduced Pheophytin in Photosystem II Discovery of the Intermediary Electron Acceptor II. Additional Demonstration of Pheophytin being the Primary Electron Acceptor of Photosystem II Interaction between and III. The EPR Spectrum of IV. Midpoint Potential of the Intermediary Electron Acceptor V. Picosecond Kinetics of Photochemical Charge Separation and Electron Transport in Photosystem II

I.

305 308 311 313 316

Oxygen Evolution 18. Oxygen Evolution

Introduction

323-336

I. Kinetics of Oxygen Evolution II. Proton Release accompanying S-State Transitions

19. Oxygen Evolution I. II. III. IV. V.

The Role of Manganese

Methods for Extracting Manganese from Photosynthetic Organelles Monitoring Manganese by EPR Spectroscopy Manganese Stoichiometry EPR Spectra of the Manganese Complex in the A Manganese-Cluster Model based on EPR and X-Ray Absorption Spectroscopy

xv

324 331

337-354 338 339 340 342 344

20. Oxygen Evolution UVAbsorbance Changes associated with S-State Transitions I. II.

355-364

Absorbance Changes produced by the Transition Absorbance Changes produced by the S-State Transition Cycle

21. Oxygen Evolution Extrinsic Polypeptides and Inorganic Ionic Cofactors

355 357

365-375

I. The Three Extrinsic Polypeptides: Properties and Functions II. Functional Role of the Inorganic Cofactors in Oxygen Evolution

22. The Electron Donor to

EPR Spectroscopy

366 369

377-396

I. The Secondary Electron Donor of Photosystem II EPR Spectroscopy II. Chemical Identity of the SII Signals III. A New Model of Oxygen Evolution Involving Hydrogen-Atom Abstraction by during the S-State Cycle

23. The Electron Donor to

OpticalSpectroscopy

I. Electron Donation from to II. Absorbance Changes and Kinetics due to

378 384 391

397-406 398 401

Oxidation by

24. Charge Recombination in Photosystem II and Thermoluminescence

407-418

I. Energetics of Thermoluminescence II. Some Commonly Observed Thermoluminescence Bands and their Origins III. Concluding Remarks

407 409 416

Photosystem I 25. Photosystem I

Introduction

419-430

I. Pigment Molecules, Cofactors, and Protein Subunits of the Photosystem-l Reaction Center II. Three-dimensional Structure of the Photosystem-l Reaction Center III. Protein Subunits of the Photosystem-l Reaction Center

26. Photosystem-l Membrane, Complexes and Crystals I. Early Work on Fractionation and Characterization of the Thylakoid Membrane II. Simplified Photosystem-l Reaction-Center Complexes III. Crystals of the Photosystem-l Reaction-Center Complex

27. Light-Harvesting Chlorophyll-Protein Complexes of Photosystem I

420 422 423

431-443 432 437 439

445-461

I.

Isolation and Spectroscopic Characterization of Photosystem-l Light-Harvesting Chlorophyll-Protein Complexes II. PS-I Core-Antenna Chlorophyll Molecules the Three-Dimensional Structure III. In Vivo Organization of Light-Harvesting Complexes in the PS-I Thylakoid Membrane

28. The Primary Electron Donor of Photosystem I I. II. III. IV. V.

Optical and EPR Properties Chemical Identity of P700 The Extinction Coefficient of P700 Redox Potential of P700 Effect of Protein Environment on P700 Properties

xvi

P700

445 451 456

463-477 464 466 469 471 473

29. The Membrane-Bound Iron-Sulfur Proteins (FeS-A and FeS-B): Secondary Electron Acceptors of Photosystem I

479-504

I.

Discovery of the Membrane-Bound Iron-Sulfur Proteins FeS-A and FeS-B and their EPR Properties II. Redox Potentials of the Membrane-Bound Iron-Sulfur Proteins FeS-A and FeS-B III. Structure and Properties of FeS-A/FeS-B (PsaC) VI. Electron-Transfer Sequence in the Reduction of FeS-A and FeS-B

480 482 483 494

30. P430: The Spectral Species Representing the Terminal Electron Acceptor of Photosystem I

505-526

I. A Brief Chronicle of the Investigation of the Photosystem-l Primary Electron Acceptor II. The Spectral Species P430: Its Discovery and Properties III. Five Possible Fates of the Photooxidized Primary Donor and the Photoreduced Primary Acceptor IV. Some Properties of P430 and P700

505 507 509 512

31. The Iron-Sulfur Center FeS-X of Photosystem I, the Photosystem-l Core Complex, and Interaction of the FeS-X Domain with FeS-A/FeS-B 527-554 I. II. III. IV. V.

FeS-X and its Spectral Properties Redox Potential of FeS-X Isolation and Characterization of the PS-I Core Complex Structure and Environment of the FeS-X Cluster in the PS-I Reaction Center Establishment of the Position of FeS-X in the Electron-Transfer Sequence

32. The Primary Electron Acceptor I. II.

of Photosystem I

Early Spectroscopic and EPR Studies of Spectra and Reaction Kinetics of by Rapid-Spectroscopic Measurements

33. The Intermediate Electron Acceptor Phylloquinone (Vitamin I. II. III. IV.

528 533 536 540 548

555-578 556 561

of Photosystem I 579-604

Occurrence of Phylloquinone in Higher Plants, Algae, and Cyanobacteria Spectroscopic Identification of a Quinone Acceptor in Photosystem I Forward Electron Transfer from to FeS-X Phylloquinone Extraction, Reconstitution, and Replacement

34. Mobile Electron Carriers Plastocyanin and Ferredoxin, and Ferredoxin Reductase I. Plastocyanin (PC) Structure and Properties II. Ferredoxin (Fd) Reductase (FNR) III. Ferredoxin

580 581 591 597

605-634 606 621 628

xvii

Proton Transport and Photophosphorylation 35. The Interphotosystem Complex

Complex and the Related 635-664

I. Topology of Cytochrome and Structure of its Subunits II. Spectroscopic Characterization of the Subunits of III. Electron Transport and Proton Translocation in Cytochrome

36. Proton Translocation and ATP Synthesis I. II. III. IV.

Introduction Composition and Structure of Chloroplast ATP Synthase The Chemiosmotic Theory and ATP Synthesis Mechanism of ATP Synthesis

Appendix: How to View the Stereograms Abbreviations Index

Complex

636 648 651

665-737 666 668 676 707

739 741 747

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Preface “Civilized man, long having abandoned his worship of fire and sun, is scarcely further advanced than his ancestors, since the fundamental relations between the sun, the green plant and himself is still shrouded in mystery. Not until the time comes when he can reproduce this apparently simple process of photosynthesis in his laboratory, independent of the green plant, can Man claim to be free from the vestiges of his ancestral worship of sun and fire. When that day comes the energy awaiting his disposal is enormous.” Pei-sung Tang Green Thralldom. London: Allen & Unwin Ltd., 1949

In the summer of 1980, I was privileged to be invited by Prof. Pei-sung Tang, then director of the Botany Institute of the Chinese Academy of Sciences, to present an extended series of lectures there on the current status of research in the field of photosynthesis. It was an enjoyable experience which allowed for coverage of biochemical and biophysical aspects of photosynthesis, in both green plants and photosynthetic bacteria. On a subsequent visit, I was urged by Prof. Tang and his colleagues at the Institute to prepare a monograph in Chinese based on the lectures I presented on the occasion of my earlier visit to the Institute. Since I was about to retire from the Charles F. Kettering Research Laboratory, I gladly accepted this suggestion and, after arming myself with many Chinese-English technical dictionaries and soliciting the able assistance of colleagues at the Institute, began work. From this effort the book “Photosynthesis: Photons, Excitons, Electrons, Protons, Ions, and Their Interactions with the Photosynthetic Membrane” eventually emerged, published by the Anhui Educational Publishing House in 1991. With few other professional obligations to be met, I was able to devote a good deal of my time to the writing of the monograph, including the preparation of all the texts and graphics in camera-ready form for publication. The present book, Photosynthesis: Photobiochemistry and Photobiophysics, evolved from that monograph, but with greater coverage and updated. This book is primarily intended for advanced undergraduates and graduate students in plant sciences, microbiology, biochemistry and biophysics. It is also hoped that the moderately detailed treatment of a limited area of photosynthesis given here may also be of some interest to workers in other, but related fields of science. Perhaps the best way to characterize the scope of the book is to note that it is concerned with photosynthetic reactions taking place in the time frame between and second, i.e., events which precede the well-studied “dark reactions,” which have been amply documented in an earlier phase of research in photosynthesis. These early reactions include a primary charge separation followed by the very rapid charge transfer leading to more stable, i.e., longer-lived species and eventually, in the case of green plants, the reduction of the terminal electron acceptor, ferredoxin, and oxidation of water. Given the very short span of time in which these rapid events occur, it is not surprising that they involve only components that are either in, or closely associated with the reaction center. In describing the scope of his wonderful little book, “Primary Processes in Photosynthesis” (1963), Martin Kamen wrote that “much of the excitement is over by the time identifiable chemical products of photosynthesis appear.” This sentiment also describes aptly the scope of the present volume. I am most grateful for the opportunity given me to prepare this book on photosynthesis, especially at a time when advances in our knowledge of the primary photochemical reactions and of the structures of reaction components has been advancing so rapidly and revealing that photosynthesis is even more clearly an integrated biological process of continuing interest and of profound importance. xix

The book is divided into five major parts: (I) An Overview, (II) Bacterial Photosynthesis, (III) Photosystem II and Oxygen Evolution, (IV) Photosystem I, and (V) Proton Transport and Photophosphorylation. Parts II, III and IV begin with an introductory chapter first, followed by chapters on the lightharvesting complexes, the primary electron donor and the primary and secondary electron acceptors, and finally the secondary donor. I have taken what might at first seem as an unnatural approach in organizing the material on the electron acceptors. I have presented the discussion of them more or less in the chronological order of their discovery rather than in the order of their actual participation in electron transfer, as dictated by their physical location or redox potential. The reduced state of an acceptor generally has a longer lifetime the further away it is from the photooxidized donor and can therefore be more readily detected with instrumentation having only modest time resolution. With the advent of instrumentation with improved time resolution, earlier acceptors with their shorter lifetimes could in turn be detected. Thus presentation in the order of discovery can better convey the sense of historical development and also illuminate some sense of the semantic difficulties that had been encountered with the term “primary.” It also underscores the importance of advances in instrumentation in realizing ever finer details of structure and mechanism which are essential to a full understanding of photosynthesis. In preparing this book, I am most grateful for being able to use results of numerous workers to illustrate the various topics. I also want to thank my colleagues for allowing me to reprocess many of the figures so that the frames and boxes, as well as the associated text, have a uniform appearance in the book. I want to thank all colleagues for granting me permission to use their figures and to allow me to make these minor modifications. I also felt very fortunate in being able to obtain a number of color photographs, particularly of the crystal structures of the various photosynthetic proteins, including those involved in electron-transport, light-harvesting and ATP synthesis. To all the contributors for their enthusiastic support, I am most grateful. I especially wish to thank Dr. Hans Deisenhofer for letting me use for the cover of this book their wonderful color picture of the crystal structure of the Rhodopsudomonas viridis reaction center. This book has used a number of stereograms showing the three-dimensional structure of various photosynthetic proteins. I especially want to encourage readers to view these fascinating pictures with the unaided eye as I am sure they will find such viewings most enjoyable and rewarding. For readers who are unfamiliar with the proper viewing procedure, a very brief instruction is presented in the appendix. At the end of each chapter there is a number of references of recent review articles as well as an extensive list of references that will provide readers access to the original work in the literature. I want to take this occasion to thank Govindjee, Chief Editor for the Advances in Photosynthesis series published by Kluwer Academic Publishers, for including this book in the series, and for his most careful and meticulous editing of the entire draft. It is also a pleasure to acknowledge the help of several colleagues and friends, Jan Amesz, Darrell Fleischman and Richard Malkin in critically examining various parts of the book and for their thoughtful and valuable comments and suggestions. I also had the pleasure and special privilege of working closely with my colleague and friend, Ed Dolan, a great grammarian, whose detailed comments on every chapter are most certainly responsible for whatever cogency the book has attained. Any omissions were due either to oversight on my part, for which I apologize, or the need to keep the book to a manageable length without distorting the major topics presented. Finally I want to thank my wife, Keiko, for her encouragement and infinite patience throughout the years that the book was being prepared. Bacon Ke 6100 Horsemans Canyon Drive, Walnut Creek, California 94595 U.S.A.; E-mail address

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Chapter 1 Photosynthesis: An Overview

I. Solar Energy and Photosynthesis II. Absorption of Light Energy and the Fate of the Electronically Excited States III. Light-Harvesting Pigment Molecules A. The Chlorophylls B. Chlorophyll Forms C. The Accessory Pigments IV. Development of Some Modern Concepts of Photosynthesis A. Van Niel s Formulation of Photosynthesis as an Oxidation-Reduction Process (1931) B. Time Sequence of Photosynthesis C. Concept of the Photosynthetic Unit” of Emerson and Arnold (1932) D. Reaction-Center and Light-Harvesting Complexes E. Discovery of the Hill Reaction (1937) V. The Thylakoid Membrane of Chloroplasts VI. The Emerson Enhancement Effect and Evidence for Two Photosystems VII. Freeze-Fracture Electron Microscopy of Thylakoid Membranes VIII. Composition of Thylakoid Membranes Lipids, Proteins, and Electron Carriers IX. Lateral and Transverse Asymmetry of Thylakoid Membranes X. Evolution of Photosynthetic Reaction Centers

1 4 6 6 8 10 13 13 14 14 16 19 19 22 26 30 38 41

I. Solar Energy and Photosynthesis Photosynthesis takes place in green plants, algae, cyanobacteria and photosynthetic bacteria where the sunlight is converted into chemical energy. In green-plants photosynthesis requires, besides light as the energy source, only two raw materials: water and carbon dioxide from the atmosphere. The organic compounds ultimately produced by photosynthesis, directly or indirectly, include sugar, carbohydrates, lipids and proteins, all of which serve as food for all living creatures. It is generally believed that the earth was formed about 4.6 billion years ago and that life on earth began about 3.5 billion years ago, with the first photosynthetic organisms to appear being the photosynthetic bacteria and primitive algae. Evidence indicates that cyanobacteria evolved about 2 billion years ago leading to the accumulation of oxygen in the atmosphere. The presence of oxygen and the production of foodstuffs by higher plants made the existence of heterotrophs such as humans possible. At present, the total biomass produced annually by plant photosynthesis amounts to about two hundred billion tons. Living creatures, including humans, consume the photosynthesized foodstuffs and derive energy from them by “respiration” (see Fig. 1), a process by which the organic compounds are oxidized back to carbon dioxide and water. Photosynthesis therefore serves as a vital link between the light energy of the sun and all living creatures.

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Fig. 1 illustrates the complementary relationship between photosynthesis and respiration. In green plants, the photophysical and photochemical processes of photosynthesis take place in the chlorophyllcontaining thylakoid membrane of the chloroplasts. The essential photochemical process includes the decomposition of water into oxygen, which is released to the atmosphere, and the generation of reducing power in the form of NADPH, plus ATP, a principal energy currency formed from ADP by a photosynthetically generated proton gradient, both of the latter two species contributing to the biosynthesis of carbohydrates and other compounds. Biosynthesis of carbohydrates from carbon dioxide occurs in the stroma region of the chloroplasts via the Calvin cycle. The formation of a six-carbon sugar molecule requires six complete turns of the Calvin cycle, for each of which three ATP and two NADPH molecules are consumed. The overall biochemistry of photosynthesis for the formation of one glucose molecule from six molecules may be written as:

Note that photosynthetic reduction of an to give an NADPH needed in this reaction two electrons, i.e., Thus the two NADPH molecules needed for the fixation of one molecule require four electrons and two protons, all originating from two water molecules. In the currently accepted scheme, two photoacts are required for the release of each electron from water, thus requiring eight moles of photons for each mole of reduced or 381 kcal. The reduction of one mole of molecule to glucose requires 112 kcal and thus the efficiency of photosynthetic reaction is 112/384, or ~30%. The actual efficiency may be lower, given that not all photons may be effective.

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Let us now examine the efficiency of solar energy utilization by photosynthesis in a broader perspective with the help of Fig. 2. This matter also concerns the problem of the earth’s energy resources, both the existing energy reserve and the continuing energy inflow from the sun. At the moment, the total fossil fuel reserve – a remnant of ancient photosynthetic products, a sort of stored solar energy bank – is driving the modern industrial civilization and is estimated to be kcal. Current world annual consumption of fuel is kW·h). So the current fuel reserve at the present rate of consumption will last about 200 years. The ultimate source of solar energy is nuclear energy, which is constantly being generated by the sun. The eventual solution for the fuel-energy needs of human kind may lie in two areas other than photosynthesis: one, sustained nuclear reaction on earth similar to that of the sun, the so-called “atomic energy,” and the other, “solar cells,” which directly convert solar energy into electrical energy. There has been slow but steady progress in the latter field. The efficiency of solar cells under development at the present time is rapidly closing in on that of conventional sources in providing cost-effective electric power. As already noted, to sustain living creatures on earth, food as well as other chemicals produced by photosynthesis are needed. Fig. 2 shows where things stand in relation to the global energy balance in the biosphere. The annual flow of sunlight energy toward the earth’s surface is estimated to be kcal Out of this a large fraction is either reflected by the atmosphere or absorbed by it . This latter portion ofenergy, although lost to photosynthesis, nevertheless powers various processes at the earth’s surface such as winds, ocean currents, hurricanes, etc. About kcal of sunlight falls onto the ocean surface, and a smaller portion amounting to kcal falls on land. Of the total, the energy stored by green plants and algae has been estimated to be kcal. From this amount, the energy from substances suitable for use as fuel amounts to kcal; foodstuff amounts to kcal, and agricultural waste amounts to kcal. Based on the total sunlight energy from the sun falling on the earth’s surface, only kcal, less than 0.1%, appears to be efficiently utilized by photosynthesis, and the energy stored in the final useful products is another order of magnitude smaller.

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II. Absorption of Light Energy and the Fate of the Electronically Excited States We briefly review here the absorption of light energy and the possible fate of the excited molecules formed. A photon is absorbed by a molecule very rapidly, within sec, provided the frequency of the incident light meets the criterion of where is the difference between the energy levels of the initial (ground) and final (excited) states, and the frequency and the wavelength, respectively, of the light, c the velocity of light, and h Planck’s constant. In most instances, the absorbed photon promotes the electron from the ground singlet state to some excited singlet state or as illustrated in Fig. 3 (A) by steps and respectively. A simplified absorption spectrum corresponding to these different changes in energy levels is illustrated in Fig. 3 (D), where the structure often observed for transitions to different vibrational levels has for simplicity been ignored. Rotational structure is usually not observed in the condensed state. A molecule brought to an upper excited state usually returns to the first excited state in sec by “internal conversion” . By this pathway a portion of the energy of the higher excited state is dissipated as heat to the surroundings by a sequence of small transitions through the vibrational sublevels of the electronically excited state.

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Although our primary interest is in the excited states produced by the absorption of electromagnetic energy, a variety of other forms of energy such as electrons and other charged particles, thermal excitation, sound and shock waves and chemical reactions, can generate excited states, and depending on the mode of excitation, the radiative decay from these excited states would be called electroluminescence, thermoluminescence, sono- or triboluminescence, and chemo- or bioluminescence, respectively. However, among all modes of excitation, that by photons is the most selective. By using light of appropriate frequency or wavelength, molecules can be transformed from a well-defined initial state to a welldefined excited state. An excited molecule in a singlet state usually has a maximum lifetime of sec due to an inherent tendency to fluoresce but can undergo various modes of nonradiative as well as radiative decay, the latter generally termed luminescence. The fate of the excited molecule depends on the pathway taken by the excitation energy. Depending on the mode of radiative decay, luminescence may also be called fluorescence, phosphorescence, after-glow or delayed light emission, etc. Once the excited molecule has returned to one of the lower vibrational sublevels of the first excited singlet state, it can return to one of the lower vibrational sublevels of the ground state by emitting a photon as fluorescence The relationship between the fluorescence spectrum and that of absorption is shown in Fig. 3 (D). Fluorescence is usually considered as a waste of the absorbed energy, but the phenomenon provides a very valuable tool for investigating photosynthesis, even though the fluorescence yield, i.e., number of photons emitted relative to the photons absorbed, is usually less than a few percent, or even lower in photosynthetic systems. Another important energy pathway is called “intersystem crossing” which refers to the conversion of an excited singlet state to an excited triplet state, followed by a radiative or nonradiative decay of the excited triplet state to the ground singlet state. The light emitted by the radiative, triplet-to-ground state transition is called “phosphorescence” The electron spins in the ground state and the excited singlet and triplet states are represented in Fig. 3 (B) and (C). Besides returning to the ground state from the state by emission, the state may alternatively go to by radiationless deactivation but from to followed by ]. As the name implies, no radiation is emitted and the electronic excitation energy is converted into vibrational excitation of which is then transferred to the adjacent solvent. For certain excited molecules the most useful decay pathway is “energy transfer” which eventually leads to the so-called charge-separated state, and is of vital importance to photosynthesis. The mechanism by which energy transfer takes place in plant pigment systems is called resonance transfer, formulated by Theodor Förster some fifty years ago. This type of energy transfer may appear like simple radiative transfer, but it is not just simply a process of one molecule emitting fluorescence and another molecule absorbing it. Instead, there is a direct coupling via the mutual radiation fields of the two oscillators, corresponding, respectively, to fluorescence in the donor molecule D and absorption in the acceptor molecule A. Resonance transfer implies a more “intimate” coupling between the energy donor and acceptor, rather resembling the interaction of weakly coupled pendulums. The oscillation associated with de-excitation of one molecule is coupled at close range to a “sympathetic” oscillation in a neighboring molecule, causing transfer of excitation energy to the latter. Thus the first molecule donates its energy directly to the second one without the intervention of fluorescence or absorption in the usual sense. Consequently, transfer is even more rapid than by radiation. It is possible for energy to be transferred by the resonance-transfer mechanism to an unexcited molecule over a distance of 50 to 100 Å. Furthermore the two molecules need not be identical. For instance, resonance energy transfer may occur from Chl b to Chl a. The one important condition for efficient resonance transfer is that the fluorescence spectrum of the donor overlap that of the absorption spectrum of the acceptor, as shown in Fig. 4.

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III. Light-Harvesting Pigment Molecules The solar energy needed for photosynthesis is captured by pigment molecules, the major pigment being the chlorophylls. There are also the so-called accessory pigments (including carotenoids and chlorophyll b where Chl a is the major pigment) that supplement the “harvesting” of light by absorbing in some wavelength region(s) in the visible where chlorophyll a itself does not absorb significantly.

II.A. The Chlorophylls Electromagnetic energy lying mainly in the visible or near-infrared region is capable of initiating photochemistry in the photosynthetic apparatus. With the principal pigment being chlorophyll (Chl). Chlorophyll contains a macrocyclic tetrapyrrole ring, with a structure similar to that of the prosthetic heme group of hemoglobin and cytochromes, and is biosynthetically derived from protoporphyrin IX. The tetrapyrrole ring of Chl [see Fig. 5 (A)] can be seen to differ from the heme in several aspects:

(1) The central metal ion in Chl is not or as in the hemes. Chl has a cyclopentanone ring (ring V) with an attached carboxylic ester group, fused to pyrrole (2) ring III. (3) Pyrrole ring IV of Chls a and b is reduced, i.e., it contain two extra hydrogen atoms. In bacteriochlorophylls a and b (BChl a and BChl b), rings II and IV are also reduced. (4) The propionyl group on ring IV of Chls a and b and BChl b is esterified by the long-chain isoprenoid alcohol phytol. In BChl a the propionyl group is esterified by either phytol or the all-trans geranylgeraniol, depending on the bacterial species [see Fig. 5 (B)].

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In addition to Chl a, photosynthetic organisms can contain other kinds of chlorophylls: Chl b in green plants and green algae, Chl c in brown algae and diatoms, and Chl d in red algae. These various chlorophylls differ by having different substituent groups on the tetrapyrrole ring, as noted in Fig. 5 (B). BChl a, present in purple bacteria, has its long-wavelength, in vivo absorption band shifted toward the near infrared (~875 nm) relative to its location in organic solvents. BChl b, found in Rhodopseudomonas viridis; has its in vivo absorption maximum at ~1020 nm and differs from BChl a in the substituent group at C-4 [see table in Fig. 5 (B)]. Another bacteriochlorophyll, called Chlorobium chlorophyll (CChl 660), is found in green sulfur bacteria, where 660 represents its long-wavelength absorption-band maximum in organic solvents; in vivo, this absorption maximum is shifted to 740 nm. In Fig. 5 (A), the conjugated systems of Fe-protoporphyrin, Chl a, and BChl a are shown shaded. The Fe-protoporphyrin IX is fully conjugated and the tetrapyrrole ring is highly symmetric. Due to the reduced state of ring IV in Chls a and b, however, the symmetry of their conjugated tetrapyrrole rings is lower. This difference in structure has a significant effect on their light-absorption properties. For instance, the long-wavelength of cytochromes, which contain the symmetrical heme, is relatively weak, whereas both Chls a and b have an intense absorption band at 676 and 642 nm, respectively [Fig. 5 (C)]. In BChl a, where the bond between C(3) and C(4) in ring II is also single, the symmetry of the conjugated system is more pronounced, resulting in an even stronger absorption band, further toward the near infrared at 780 nm. The transitions involved in the absorption and emission of light by chlorophylls and the associated spectra may be illustrated for BChl a, as shown in Fig. 5 (D). Absorption of light by BChl a, as governed by the relationship, promotes an electrons from a ground-state orbital to one of the various excited-state orbitals. In Fig. 5 (D, a), is the energy level of the molecule in the ground state, and etc. represent the energy levels of various excited states resulting from the absorption of photons of energies etc. Although the figure is scaled for the absorption spectrum of BChl a, qualitatively it is equally applicable to Chl a or Chl b. In Fig. 5 (D, b), the absorption spectrum is shown turned 90° clockwise from its conventional orientation in order to relate it to the energy diagram. The process of light absorption by Chl may be written as In chlorophyll, the optical transition axes x and y lie in the plane of the system of the tetrapyrrole ring along each of the N-N diagonals [see Fig. 5 (A)]. The absorption spectra of chlorophylls in solution show two main pairs of bands, one pair in the blue region designated as the B-bands) and one pair in the red or far red designated as the Q-bands). The subscripts x and y are used in Fig. 5 (D, b) as they were in Fig. 5 (A). Following the discussion given in section II, above, we see in Fig. 5 (D, c) that the upper excited states of chlorophyll undergo extremely rapid radiationless deactivation to the lowest excited state. The lowest excited state of chlorophyll in solution has a maximum lifetime of ~5 ns, it decaying to the ground singlet state, in part by fluorescence, with the corresponding spectrum shown in Fig. 5 (D, d). Some portion of the population of the lower singlet excited states may also undergo intersystem crossing to the lowest triplet state as shown earlier in Fig. 3,

lll. B. Chlorophyll Forms Whereas pure chlorophyll in dilute solution displays a simple absorption spectrum with relatively narrow bandwidths, the absorption bands of chlorophyll in vivo are usually displaced to longer wavelengths as a result of being bound to protein and the bands are also much broader. More than sixty years ago Albers and Knorr1 (of the then “C. F. Kettering Foundation for the Study of Chlorophyll and Photo-

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synthesis”) measured the in vivo absorption spectrum of chlorophyll in individual chloroplasts of three algae by means of a microspectrophotometer specially constructed from a microscope and a high-resolution spectrograph using photographic plates calibrated for intensity and wavelength. The in vivo absorption band between 664 and 704 nm generally had a broader bandwidth than the corresponding band in solution, A more striking feature seen in the spectra of all chloroplasts examined was the presence of several absorption maxima. The relative intensities of the maxima varied from a cell of one species to that of another, indicating that they are due to not one but several different substances. For instance, in the alga Protococcus, bands with absorption maxima could be identified at 673, 681, 687 and 698 nm each with an intensity characteristic of the alga. Albers and Knorr suggested that in vivo chlorophyll a most likely formed “reaction products” of some kind having absorption bands at slightly different wavelengths. In the late 1950s, Stacy French2 renewed the concept of “ chlorophyll forms.” The availability of more advanced spectrophotometer designs and newer methods for data manipulation helped expedite further studies. In addition, workers around that time were beginning to prepare subchloroplast fractions having different pigment compositions, different Chl a-to-Chl b ratios and therefore different absorption spectra. These subchloroplast fractions isolated from both plants and algae provided useful experimental materials for studying the various forms of chlorophyll. At the same time French3 successfully applied curve-fitting techniques and pioneered the use ofautomatic recording of derivative spectra for quantitative analysis as well as identification of chlorophyll forms. The existence of different chlorophyll forms in vivo can best be seen by comparing the absorption spectrum of pure chlorophyll a in ether with the spectra of two algae that are known from column separation work to contain Chl a only. As many studies had already shown that both the band shapes and peak positions of in vivo Chl a varied in the many algae and plants that had been examined, it is not surprising that the in vivo absorption spectra of Cryptomonas and Botryoglossum, seen in Fig. 6 (A), had the absorption band red-shifted from solution spectra, but the spectra also indicated the presence of Chl a in different “forms” in the two algae. Different chlorophyll forms of Chl a had also been detected in a single organism, as illustrated in Fig. 6 (B), where the absorption spectrum for a preparation of the alga Euglena is compared with that of pure Chl a in ether. We see that the native Chl a in its functional, in vivo state has a much more complex spectrum consisting of a number of overlapping absorption peaks. From this absorption spectrum, four Chl-a forms were distinguished and designated as Ca 672, Ca 681, Ca 693 and Ca 706, where the numbers represent peak wavelengths. Fig. 6 (C) illustrates the use of curve-fitting as a means of characterizing the various Chl forms in Scenedesmus fraction I (i.e., a so-called “photosystem-I” preparation). Here the observed spectrum, plotted as dots, was resolved by computer into a number of Gaussian-shaped distribution curves of different heights, widths and location, each curve representing different chlorophyll forms. For instance, curve analysis of the 653-nm Chl-b band (not shown) yielded a major 654-nm form and a minor 645-nm form. As for Chl-a, the major forms are at 662, 670, 677 and 683 nm, plus two minor forms at longer wavelengths, namely, at 693 and 702-703 nm. The four major forms just mentioned were considered the “universal” forms of Chl a. It has been shown that taken together these four major components plus the two longer wavelength components can give a good fit to many spectra. As illustrated in Fig. 6 (C), adding together the component spectra gave a spectrum (the solid curve) which runs almost exactly through the experimental data (the dots). The peak wavelengths and their approximate distribution in the two fractions from green algae and leaves are listed in the table next to Fig. 6 (C).

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III.C. The Accessory Pigments Besides the major light-absorbing chlorophyll molecules, there are two groups of accessory pigments which absorb light in the wavelength region where chlorophylls do not absorb strongly. The two types of accessory pigments are the carotenoids and phycobilins. More than 600 naturally occurring carotenoids are known at the present time, of which about 150 are known to be present in photosynthetic organisms, including higher plants, algae and photosynthetic bacteria. Carotenoids are long-chain, conjugated hydrocarbons containing a string of isoprene residues,

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and distinguished from one another by their end groups. Usually, a few different carotenoids are present in each organism. Fig. 7 shows four carotenoids lutein, violaxanthin and neoxanthin) commonly present in green plants and spheroidene and the carotenoid commonly present in photosynthetic bacteria, together with their full names and the wavelengths of their three absorption maxima in the visible region. Note that is one of the three common end groups, and (out of a total of seven), present in green-plant and algal carotenoids. Carotenoid absorbance extends from the UV to about 550 nm in the visible, as illustrated by the characteristic three-peaked spectrum of shown in Fig. 9 below. The short wavelength peak in a carotenoid is usually attenuated if there is a group present. In any event, the characteristic absorption spectrum of carotenoids is responsible for the yellow color of leaves in autumn and the orange color of carrots, for which this group of pigments is named. The upper wavelength limit of absorption by carotenoids depends not only on the number of conjugated double bonds and the isomeric configuration, but also on the nature of the end groups, in particular the presence or absence of oxygen (as epoxides) or hydroxyl groups.

Invariably carotenoids are present together with chlorophylls in photosynthetic organisms, where carotenoids serve a dual function. The first role is as an accessory light-harvesting pigment. The efficiency of singlet-singlet energy transfer from certain carotenoids to chlorophyll may be very high, ranging from 70% to nearly 100%. The other role of carotenoids is as a “quencher” of triplet chlorophyll to provide protection against photo-oxidative damage. In fact, carotenoid-deficient mutants of algae or photosynthetic bacteria are easily destroyed by the combined presence of light and oxygen. In addition to the two well-established roles, recent reports have suggested a structural role for some carotenoids in green plants. Havaux4 reported that carotenoids of the xanthophyll family and some other terpenoids may stabilize and photoprotect the lipid phase of thylakoid membranes. Available evidence

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suggests that, when plants are exposed to potentially harmful environmental conditions such as strong light and/or elevated temperatures, the xanthophyll, violaxanthin, and the products of their enzymatic de-epoxidation may become partitioned between the light-harvesting complexes and the lipid phase of the thylakoid membranes. The resulting interaction between the xanthophyll molecules and the membrane lipids brings about a decrease in membrane fluidity, an increase in membrane thermostability and a lowered susceptibility to lipid peroxidation. The second group of non-chlorophyll, light-harvesting pigments are the phycobilins, which are abundant in red algae and cyanobacteria (formerly called blue-green algae). Phycobilins are responsible for these organisms’ distinctive colors, after which they are named. Phycobilins are non-cyclic tetrapyrroles, i.e., structures that may be considered as a chlorophyll tetrapyrrole ring opened up and the Mg atom discarded. There are four different phycobilin pigments present in the phycobiliproteins of cyanobacteria and red algae. Fig. 8 shows how two major phycobilins, phycocyanobilin and phycoerythrobilin are linked to polypeptides to form phycobiliproteins. Phycocyanobilin is linked to a polypeptide with its ring A through a thioether linkage (top row). Phycoerythrobilin is similarly linked, either singly with ring A or doubly with rings A and D, to one or two polypeptides through thioether linkages as shown in the bottom row of Fig. 8. Structural differences arising from these bilin-to-polypeptide linkages, the amino-acid sequence of the polypeptide and the chemical composition at the attachment site are responsible for the complex and varied absorption spectra of the phycobiliproteins in vivo. In cyanobacteria and red algae, the phycobiliproteins are assembled into supramolecular aggregates called phycobilisomes, which are appressed to the photosynthetic membrane surface and transfer electronic excitation energy to the reaction centers via chlorophyll to initiate photochemical reactions.

Carotenoids and phycobiliproteins, as is made clear by their designation as accessory pigments, serve to absorb light energy not absorbed in the spectral region absorbed by chlorophyll and transfer this energy to chlorophylls. Thus chlorophylls complemented by the accessory pigments, as shown in Fig. 9, can cover more fully the solar spectrum from the near ultraviolet to the near infrared and thus achieve a more efficient utilization of solar energy. The light-harvesting chlorophyll- and bacteriochlorophyll-protein complexes will be discussed in detail in chapters 3, 12, and 27. The carotenoids, their light-harvesting role, the energy-transfer dynamics involved in light harvesting, and the structure and properties of a supramolecular chlorophyll-carotenoid (fucoxanthin) complex – a “xanthosome” – will be presented in chapter 13. The phycobiliproteins and the supramolecular phycobilisomes will be presented in detail in chapter 14.

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IV. Development of Some Modern Concepts of Photosynthesis

IV.A. Van Niel s Formulation of Photosynthesis as an Oxidation-Reduction Process (1931) During the early part of this century, Richard Willstätter and Arthur Stoll5 along with Otto Warburg6 believed the function of light is to activate and decompose the molecule into carbon and oxygen. The carbon part then believed to combine with to form carbohydrate. In this hypothesis, the released oxygen originated from However, it was known that photosynthetic bacteria can assimilate without evolving oxygen and, furthermore, bacterial photosynthesis requires the presence of a suitable reducing substrate (a generic designation, where A in the case of photosynthetic bacteria is not oxygen). For photosynthetic bacteria, the reducing substrate may be various simple organic 7,8 compounds or simply itself. Around 1930, Cornelis van Niel thus formulated the process of bacterial photosynthesis process as follows:

where represents carbohydrate formed by photosynthesis. Van Niel then considered photosynthesis in green plants and algae a variation of this process, where the reducing substrate is and thus formulated the reaction as:

For both cases, van Niel was able to express his postulated reaction in rather formal terms as the production of an oxidant “S” or “OH” and a reductant “H.” In other words, light produces a coordinated oxida-

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tion-reduction reaction. Or, in a more modern expression, as follows:

Accordingly, in the current view, the absorption of light is understood to produce a primary photochemical charge separation, between a chlorophyll-type, primary electron donor and a primary electron acceptor, resulting in the reactions going through a series of intermediate electron-transfer steps. One important consequence of van Niel’s formulation is that in green-plant photosynthesis, the evolved should originate from not This was soon confirmed experimentally by Ruben, Randall, Kamen and Hyde9 by mass-spectrometric examination of oxygen evolved by the alga Chlorella suspended in water. In retrospect, van Niel’s simple and elegant postulate had provided one of the most important cornerstones for our understanding of the mechanism of photosynthesis during the past half century. For an in-depth discussion of the evolution of van Niel’s concept, see the review by

IV.B. Time Sequence of Photosynthesis Time is a very special parameter in the molecular processes of photosynthesis. From the initial absorption of a photon to the beginning of biosynthesis the span of time is well over ten orders of magnitude. As shown in Fig. 10, the sequence of photosynthesis begins with an extremely fast photophysical act, followed by rapid photochemical processes, electron-transfer reactions, then relatively slower biochemical reactions, and finally still slower physiological reactions. For events taking place over a wide range of “eras”, Kamen10 introduced a logarithmic time scale called which is analogous to the expression of pH for concentration. For instance, for the sec that it takes for electronic excitation, would be the logarithm of the reciprocal of time (in seconds), namely, 15. Kamen’s logarithmic time scale offers us some perspective of the fact that different processes in photosynthesis take place in characteristic eras covering a wide time span. The extremely complex processes involved in photosynthesis and the characteristic time demarcation for each stage are indeed nature’s most amazing phenomenon. Kamen expressed it well in the introduction to his book Primary Processes in Photosynthesis (1963, Academic Press) by noting that “Few phenomena in natural science equal photosynthesis in sweep and grandeur.”

IV.C. Concept of the Photosynthetic Unit of Emerson and Arnold (1932) In green-plant photosynthesis, chlorophyll molecules, the light-harvesting antenna, absorb photons and transfer the electronic excitation energy to the reaction center, which then initiates the photochemical charge separation [see Fig. 11 (A)]. Two chlorophyll molecules, in particular, the special pair, constitute the primary electron donor in the reaction center, while the bulk of the chlorophyll, the lightharvesting molecules, serves only to capture the energy of a photon and funnel it to the special pair. In 1932 Robert Emerson and William Arnold11 investigated the question of quantum requirement for the photosynthetic production of oxygen from water. The authors illuminated a Chlorella suspension with trains of flashes with different intervals of darkness between flashes for different trains, thus allowing for subsequent dark reactions to occur following each flash. Because of the short duration of the flash, the photosynthetic reaction center can only turn over once during each flash excitation. They

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then measured the amount of oxygen evolved per flash as a function ofthe dark interval between successive flashes. It was found that the flash yield of oxygen evolution was dependent on the duration of the dark interval, with the oxygen yield increasing with increasing dark intervals up to ~40 ms, where it reached a maximum and then remained unchanged for dark intervals beyond 40-ms. These results implied that photosynthesis consists of light and dark reactions, which act together to utilize the energy of absorbed photons to the maximum extent. In this case, the dark interval is considered to provide the needed time for the chemical reactions involved in oxygen evolution to come to completion before the next flash can be fully effective. With the dark interval maintained at Emerson and Arnold then examined the oxygen yield as a function of the flash intensity. Starting with low, non-saturating, flash intensities, the yield of oxygen release increased with intensity until it reached a maximum level for saturating intensities. With flashes of saturating intensity, the maximum yield per flash based on the number of chlorophyll molecules was 1 per ~2400 chlorophyll molecules. Since 8 quanta absorbed by chlorophyll can yield one the question arose as to why the maximum yield was not 1 per 8 chlorophylls or even higher? Emerson and Arnold12 explained that 2400 chlorophyll molecules act as a unit, as illustrated in Fig. 11 (A), for producing one per 8 photons absorbed. The collective of antenna chlorophylls plus the molecular machinery for carrying out chemical reactions in the dark was called the “photosynthetic unit.” Eight photons absorbed by the 2400-chlorophyll collective would provide the necessary energy for evolving 1 molecule. The energy of any extra photons absorbed by the collective during the short flash could not be used to promote photosynthesis but only be converted to thermal energy or re-emitted as fluorescence and, in any case, wasted. Note also that a long-standing controversy existed between Otto Warburg and Robert Emerson on the question of quantum yield of photosynthesis in green plants, on the respective values of 3-4 vs. 8. See Govindjee’s fascinating paper13 for a historical perspective on this matter.

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Since one photon can initiate an electron transfer, one might surmise that there are eight cooperatives, each handling one photon, and each containing ~300 chlorophyll molecules (2400/8), and together supplying the necessary energy for the chemical processes. The energy diagram in Fig. 11 (B) illustrates exciton transfer among the light-harvesting chlorophyll molecules and the eventual trapping by the reaction-center chlorophyll. The size of photosynthetic units vary among different species: photosynthetic bacteria contain similar but much smaller photosynthetic unit of ~60 BChl-a molecules, while the green bacteria, on the other hand, have very large photosynthetic units containing ~1000 BChl-c molecules.

IV. D. Reaction-Center and Light-Harvesting Complexes In all natural photosynthetic systems, pigment molecules are bound to polypeptides forming pigmentprotein complexes. All photosystems are built on a general theme but also with some variations. All photosystems are composed of three major components: the reaction center that carries out photochemical charge separation and electron transport, the core antenna or inner antenna consisting of pigment proteins that are an integral part of the reaction center complex, and the peripheral or outer antenna. The energy-transfer pathway is from the peripheral antenna complex to the inner or core antennae and finally to the reaction center. In photosystem I of green plants, for example, there are ~90 core-antenna chlorophyll-a molecules bound to the two major protein subunits of the reaction center (for details see Chapter 27). Models of various photosystems based on this common theme with several variations are presented in Fig. 12. The purpose of the variety and complicated organization of the light-harvesting systems is to achieve an efficient collection and transfer of light energy under different ambient light conditions. Chl a is present in all organisms that have been studied and that are capable of oxygenic photosynthesis. It is present in both the light-harvesting complexes and the reaction centers as well. In the thylakoid membrane of green plants Chl a is found to be localized and at a very high local concentrations. A pigment closely related to Chl a has been implicated as the constituent of the primary donor P700 of photosystem I of green plants (see Chapter 28). Chl a is also considered to act as the first electron acceptor (called in a series of electron acceptors in the PS-I reaction center [see Chapter 32]. Similarly Chl

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a and BChl a serve as the primary donors in the photosystem II and the bacterial reaction centers, respectively. Pheophytins, i.e., chlorophylls whose Mg is replaced by two hydrogen atoms, are known to play important roles as intermediary electron acceptors in both green plants and photosynthetic bacteria. In those photosynthetic purple bacteria which contain bacteriochlorophyll a there are two spectrally different forms of the light-harvesting bacteriochlorophyll-protein complex: the core antenna, called the B890 complex, and the peripheral antenna, called the B800-850 complex [see Fig. 12 (A)]. Usually there is a fixed stoichiometry between the B890 complex and the reaction-center, but the number of B800-850 complexes may vary according to the growth conditions, such as light intensity. In bacteria containing bacteriochlorophyll b, such as Rhodopseudomonas viridis, there is a similar architecture. Their respective antenna, however, absorb further into the near-infrared. The bacterial light-harvesting complexes are discussed in detail in Chapter 3. In the green photosynthetic bacteria the bulk of the light-harvesting pigments are located predominantly in supramolecular structures called chlorosomes which are not embedded in the membrane, but, instead, are appressed to the cytoplasmic membrane through a crystalline baseplate protein, as shown in Fig. 12 (B), left, for Chloroflexus aurantiacus. In the green sulfur bacteria Chlorobium, an additional sub-antenna complex in the form of a two-dimensional crystalline array of trimeric BChl a-protein (now called the “FMO protein,” for Fenna, Mathews and Olson) is present at the attachment site between the baseplate and the chlorosome, as shown in Fig. 12 (B), right. In fact, the first crystal structure determination of any photosynthetic pigment-protein was obtained for this BChl a-protein nearly 25 years ago by Fenna, Mathews, Olson and Shaw13 at a resolution of 2.8 Å and later refined to Typically, each chlorosome contains as many as 10,000 BChl-c molecules, some carotenoids and several hundred BChl-a molecules serving as the sub-antenna in the baseplate and as core antennae in the cytoplasm. Chlorosomes and the BChl a-protein, or the FMO protein, are discussed in Chapter 8. The cyanobacteria and red algae [Fig. 12 (C)] contain chlorophyll a but no chlorophyll b. However, they contain phycobilin pigments in the form of phycobiliproteins which are aggregated into supramolecular complexes called phycobilisomes. Phycobilisomes, like chlorosomes, are also appressed to the stromal surface of the photosynthetic membrane and serve to absorb and transfer light energy primarily to the core antenna of the PS-II reaction center [see Fig. 12 (C)]. Again, the primary function of this unique light-harvesting apparatus is to allow the organism to survive under the generally weak light condition found in its environments. In view of the fact that it is mainly green and yellow light that is attenuated down through the depth of water overlying these organisms, phycobilisomes possess appropriate absorption characteristics to help absorb photons in these particular spectral regions. Phycobilisomes can funnel the absorbed energy to the reaction center of photosystem II in cyanobacteria and red algae with more than 95% efficiency. The structure and energy-transfer properties of phycobilisomes are discussed in detail in Chapter 14. Recently, a supramolecular complex consisting of chlorophyll and carotenoid has been discovered by Tetsuya Katoh16 and coworkers in the membrane of the photosynthetic brown algae Dictyota dichotoma and Petaloniafascia. This 520-kDa, orange-brown complex is made up of seven protein subunits, each with a molecular mass of 74 kDa and each containing 13 molecules of chlorophyll a, 3 Chl c, 10 fucoxanthin and 1 violaxanthin. This complex has an unusually high pigment content, with a pigment-toprotein ratio of 0.37 by weight. Katoh named the complex “fucoxanthin-chlorophyll-protein assembly,” abbreviated as “FCPA” [see Fig. 12 (D)]. More recently, this and other chlorophyll-carotenoid supramolecular complexes have acquired a new name of “ xanthosotnes.”17 The spectral properties of FCPA and arrangement and interactions of the pigment molecules in the complex are discussed in detail in Chapter 13.

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Green plants contain two photosystems designated II and I, each of which is composed of reaction centers plus core and peripheral antennae [see Fig. 12 (C), (D) and (E)]. Photosystem II contains the peripheral complex called “LHC II,” which was discovered in 1966 by Ogawa, Obata and Shibata18 during their fractionation of chloroplast thylakoids by SDS-PAGE (sodium dodecyl-polyacrylamide gel electrophoresis). There are four different LHC IIs, designated as LHC IIa through LHC IId. One of these, the LHC IIb complex, accounts for 2/3 of all the chlorophyll in the PS-II antenna, and is in fact the most abundant membrane protein of the thylakoid membrane. As a peripheral antenna, LHC IIb transfers energy to the two inner antennae called CP47 (chlorophyll protein, with molecular mass of 47 kDa) and CP43, which in turn transfer energy to the reaction center. Photosystem I contains fewer lightharvesting complexes than photosystem II, but its core antenna has nearly 100 Chl-a molecules bound to the two major protein subunits of the PS-I reaction center. The LHC I complex, present in two forms (LHC Ia and LHC Ib; see Table 3 on p. 33), has a Chl a/b ratio of 3-4, somewhat higher than LHC II. The PS-II and PS-I light-harvesting complexes are discussed in detail in Chapters 12 and 27, respectively.

IV. E. Discovery of the Hill Reaction (1937) In 1937, Robin Hill 19 succeeded in isolating chloroplasts from leaves which under suitable conditions still retained some form of photosynthetic activity. The material isolated by Hill actually consisted of thylakoid membranes which lacked the necessary enzymes for fixation as a result of the loss of the outer membrane envelope. When these “chloroplasts” were illuminated, no oxygen evolution could be observed. However, when an electron acceptor such as ferric oxalate, as was used by Hill initially, was added, oxygen evolution was observed upon illumination, accompanied by the reduction of the ferric oxalate to the ferrous form. Furthermore, four equivalents of the oxidant were found to be photochemically reduced for each mole of oxygen evolved. This reaction was called the “Hill reaction.” Later, ferricyanide and benzoquinone were commonly used as the oxidizing agents. At this juncture, van Niel’s concept of a redox reaction taking place in the reaction center became very useful for understanding the Hill reaction. Although was not assimilated under illumination, it appeared that the other reactions responsible for the splitting of water were still active. The ferric oxalate used by Hill apparently served as a substitute for the natural oxidant to intercept electrons produced at the reaction center and thus allow oxygen to be evolved. The Hill reaction shows that assimilation and oxygen evolution are not obligatorily linked and two physically distinct enzymatic systems may exist.

V. The Thylakoid Membrane of Chloroplasts Previously, we have described the molecular structure of several kinds of light-absorbing pigment molecules and the possible fate of various excited states of these molecules after light absorption. For photosynthesis to proceed, the excitation energy must be transferred, preferably with high efficiency, from the light-harvesting pigment molecules to the photosynthetic reaction center, where photochemical charge separation can then take place. We now examine the structure of the photosynthetic apparatus where all the important events leading to photochemical charge separation in the reaction center and subsequent electron transfers take place. In green plants, chloroplasts are the site of photosynthesis, and we begin by examining their presence, location and structure in leaf tissue at several different organizational levels, starting with the leaf itself, as illustrated in Fig. 13. Figs. 13 (A) and (B) are sketches of a (spinach) leaf and a magnified cross-

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sectional view of the leaf, respectively. The chloroplasts, shown in Fig. 13 (B) as gray spots around the cell perimeter, are easily seen with a low-power microscope, their being several microns in size. In green plants, there may be several hundred chloroplasts in each cell, taking up as much as 25% of the total cell volume. Fig. 13 (C) shows an electron micrograph of a single, ellipsoid-shaped chloroplast, accompanied by a sketch in Fig. 13 (C') depicting its inferred three-dimensional structure. Each chloroplast is bounded by an envelope of a highly permeable outer membrane and a nearly impermeable inner membrane, the two membranes being separated by a narrow, inter-membrane compartment [see Fig. 13 (C) and (C')]. The outer membrane allows small molecules to pass through, while the inner membrane presents a barrier which allows only certain metabolites or ATP to pass through with the help of special transport proteins embedded in the membrane. Enclosed by the inner membrane is the “stroma,” a concentrated solution containing the enzymes necessary for fixation, i.e., its conversion into carbohydrates. The stroma also contains the chloroplast’s own DNA, RNA and ribosomes involved in the synthesis of proteins. This chloroplast stroma is analogous to the matrix in mitochondria. Suspended in the stroma is a third kind of membrane, or rather, an extensive internal membrane network, called the “thylakoid membrane.” The term “thylakoid,” first coined by Wilhelm Menke20 in 1962, means “sac-like.” Note that the thylakoid membrane, unlike a similar membrane in mitochondria, is repeatedly folded back on itself into double membranes. In certain regions of the membrane network are stacks of flattened disk-like sacs called “grana” (singular, granum), each chloroplast contains about 10 to 100 such grana. The grana are interconnected by the unstacked “stroma lamellae” (singular, lamella), forming a single continuous membrane network suspended in the stroma. For better visualization of the thylakoid-membrane structure, a portion of Fig. 13 (C) is further magnified in Fig. 13 (D) and a sketch in Fig. 13 (D') shows in simplified form a portion of the thylakoid membranes illustrated in Fig. 13 (C'), where some useful terminology is also presented. In Fig. 13 (D) is a magnified view of part of (C), where the few dark spots are the osmiophilic lipid granules. Fig. 13 (D') also introduces some other terms for the thylakoid structure: the appressed region, i.e., the closely adjoined surfaces between the granal sacs, the non-appressedregions of the thylakoids that face the stroma either in the lamellar region or at the end region of the grana, and the intra-thylakoid space, the latter is also commonly called the “lumen.” Since all grana are interconnected by the unstacked stroma lamellae, the lumen of each thylakoid region is also connected with the lumen of all other thylakoid regions. In summary, chloroplasts may be viewed as consisting of three separate compartments confined by membranes: first the inter-membrane compartment between the outer and inner envelope membranes; second the stroma enclosed by the double envelope membrane and the inner membrane, and finally the compartment enclosed by the thylakoid membrane network. The thylakoids and stroma are the sites of the so-called light and dark reactions of photosynthesis, respectively. This compartmentalization of photosynthetic functions was recognized by Park and Pon 21 when they broke open the chloroplasts, separated the contents into thylakoid and stroma fractions and examined their properties. The specific activities of the thylakoids include photochemical reactions, electron transport, oxygen evolution, ATP synthesis and reduction, while the stroma contains enzymes for fixation driven by ATP and NADPH and other biochemical reactions in the dark. Our understanding and appreciation of the detailed structure and organization of the thylakoid membranes has increased tremendously in recent years. Further discussion of thylakoid structure will be continued in section VII on page 26.

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VI. The Emerson Enhancement Effect and Evidence for Two Photosystems For a given photosynthetic process, for instance, oxygen evolution, one may express its efficiency in terms of a “quantum requirement,” i.e., number of photons needed for each oxygen molecule produced or, more commonly, in terms of its reciprocal, the “quantum yield,” i.e., number of oxygen molecules produced per photon absorbed. When quantum yield is measured as a function of the wavelength of light used, then one obtains the so-called “action spectrum” for oxygen evolution. In 1943, Robert Emerson and Charlton Lewis22 examined the action spectrum in the visible region for oxygen evolution in the green alga Chlorella pyrenoidosa and found the quantum yield remained fairly constant up to 685 nm, beyond which it declined sharply, even though there is appreciable chlorophyll absorption up to 700 nm. This phenomenon, called the “red drop” by Emerson, was also observed later by Tanada23 in the diatom Navicula minima and by Haxo and Blinks 24 in green and brown algae. The red drop phenomenon could not be easily explained since photosynthesis would be expected to be brought about by the lowest vibrational state of the first excited singlet state of chlorophyll a, and absorption anywhere within the red band should very rapidly lead to this state. In 1957 Emerson, Chalmers and Cederstrand25 found, however, that the yield of photosynthesis in farred light beyond ~690 nm could be increased dramatically by supplementary light of somewhat shorter wavelength, e.g., at 650 nm, as shown in Fig. 14 for Chlorella. With supplementary light, the yield was approximately constant out to longer wavelengths before turning downward. The wavelength where the yield declines to one-half the maximum level corresponds to 718 nm in the presence of supplementary light, vs. 690 nm in the absence of supplementary light. More significantly, the quantum yield obtained by illuminating with light of two wavelengths together was actually higher than the sum of the quantum yields obtained when light of either wavelength was applied separately. This important finding is called the “Emerson enhancement effect.” Since the wavelength of the supplementary light must be of a shorter wavelength than the wavelength of any light which by itself gives a diminished quantum yield, the significance of the supplementary light could be that the maintenance of maximum efficiency of photosynthesis requires the excitation of two different pigments or pigment systems, each with a different wavelength requirement for excitation and the two systems linked in series.

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In 1960, Robin Hill and Fay Bendall 26 first attempted to summarize the various observations discussed above by a provisional “zig-zag” scheme based on a redox-potential diagram, as shown in Fig. 15 (A). The scheme proposes two photoreactions in series involving two components, X and Y. At the end of one photosystem is the reduction product XH and at the other end an oxidation product YOH. The two components, X and Y, whose nature remained obscure at the time, were initially thought to be some unspecified forms of chlorophyll. Cytochromes f and both of which were discovered byHill 27,28 , were proposed to act as carriers of reductive power (electrons) in the dark reaction that connects the two photosystems as shown by the two-light reaction scheme at the bottom of Fig. 15 (A). It was also postulated that this reaction is coupled to phosphorylation, a reaction converting ADP to ATP. The normally oxidized would have to be reduced by Y to produce YOH and by X to produce XH. Each cytochrome was known to be present in chloroplasts at a concentration of 1 per ~300 Chls, which is consistent with that expected from the concept of photosynthetic unit containing a large number of antenna chlorophyll. In 1965 Hill 29 elaborated the two-photosystem scheme further as shown in Fig. 15 (B). In this Zshaped scheme, two groups of chloroplast components with known redox potentials were placed at the bends of the “Z”: Cyt f, plastocyanin and P700, close to +0.4 V, and plastoquinone and close to 0 V. Ferredoxin, with a potential of -0.43 V, is close to the midpoint potential of hydrogen electrode. For oxygen production, the midpoint potential of the unknown component must exceed that of the oxygen electrode. Over the past thirty years, a variety of Z-schemes have been published in the literature to illustrate the electron-transfer processes in green-plant photosynthesis, but their basic features have not deviated from that shown in Fig. 15 (B). For instance, we show a currently accepted, concise Z-scheme in Fig. 15 (C); it includes many more individual components than were originally envisioned, plus a representation of the operation of the so-called Q-cycle in the To explain the Emerson enhancement effect, Govindjee and Rabinowitch 30 obtained evidence from the action spectra of the enhancement effect the presence of two distinct forms of Chl-a in vivo presumably present in two different pigment systems. Soon, new experimental evidence in support of the two-photosystem concept was reported independently by Govindjee et al. 31 and by Kautsky et al.32 from evidence obtained from fluorescence and by Kok and Hoch33, Duysens 34 and Witt35 based on measurements of absorbance changes. We present here two results based on absorbance changes associated with changes in the redox state of electron carriers in the chloroplast. In 1961, Duysens, Amesz and Kamp34 studied the two-light effect on the red alga Porphyridium cruentum by monitoring changes of the absorption spectrum that accompany the redox change of cytochrome f For excitation of the two photosystems, they used 680 nm and 560 nm lights absorbed predominantly by Chl a and by phycoerythrin of photosystems I and II, respectively. The two excitation lights were used either separately or together. As seen in Fig. 16 (A), 680-nm light alone caused transmission increase at 420 nm, indicating oxidation of Cyt f. Turning the 560-nm light also on at the same time caused a transmission decrease, indicating Cyt f reduction. The simplest interpretation of these results is that there are two pigment systems that respond unequally to light at the two different wavelengths and which interact with cytochrome f in an opposing manner. In the presence of DCMU [3-(3,4-dichlorophenyl)1, 1-dimethylurea], a herbicide inhibitor for oxygen evolution, the reduction of oxidized Cyt f by 560-nm light was blocked [Fig. 16 (B)]. In fact, when photosystem II was inhibited by DCMU, the 560-nm light actually had a small effect in promoting Cyt f oxidation. In the meantime, Witt and colleagues35,36 observed similar effects on the redox state of cytochrome f when red and far-red light were alternately used for illumination. They also observed similar two-light effects with respect to a species that was later identified as plastoquinone, as well as another species, “P700” discovered earlier by Kok. Fig. 17 (A) shows the spectroscopic results of the two-light effect on P700 in broken spinach chloroplasts. Illumination of chloroplasts with 720-nm light of 0.5-sec duration

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elicited an absorption decrease at 704 nm, representing photooxidation of P700. After excitation with far-red light, P700 remained for the most part oxidized, as indicated in the trace. A 638-nm flash of 0.04sec duration was then applied, causing a rapid absorption increase and a complete reversal of the original absorption change, indicating that the 638-nm light activated photosystem II and generated a reductant for The two-light effect or the two-photosystem concept was further supported by the action spectrum shown in Fig. 17 (B) for the redox reaction of P700 monitored at 433 nm (instead of 704 nm). The results are consistent with the notion that P700 photooxidation occurred as a result of light-induced, charge transfer in photosystem I, while the reductant for was generated by photosystem II. From the slight difference in the profiles of the two action spectra, it was surmised that longer-wavelength absorbing forms of Chl a-682 and -695 were present in photosystem I while Chl a-650 and Chl a-678 could be assigned to photosystem II.

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Rumberg et al.37 measured an absorption change due to plastoquinone that is known to be located on the reducing side of the PS-II reaction center. An absorption decrease at 254 nm attributable to the reduction of plastoquinone could be induced by a 638-nm flash. This photoreduced plastoquinone could then be completely reoxidized by far-red light at 720 nm. It was also noted that the electron-transfer reactions in the re-reduction of and re-oxidation of reduced PQ were kinetically coupled.

VII. Freeze-Fracture Electron Microscopy of Thylakoid Membranes In green plants, light absorption and electron transport take place in chloroplasts in two regions of the thylakoid membrane, in the grana consisting of stacked thylakoid membranes and in the stromal lamellae, as shown previously in Fig. 13 (C) and (D). The thylakoid network suspended in the stroma may be looked upon as composed of the intra-thylakoid space, or enclosed lumen by the grana, and the interconnecting stromal lamellae. Structural details of thylakoids have been obtained primarily by electron microscopy, using the freeze-etch and freeze-fracture technique. We first describe briefly the procedure and terminology used in the freeze-etch technique, viewed from the results obtained. Consider a small section of the thylakoid membrane in Fig. 18 (A) and its representation in the sketch in Fig. 18 (B), top. Consider also that the thylakoid specimen has been quick-frozen by immersion in freon cooled in liquid nitrogen. By sublimation of the ice surrounding the membrane in a vacuum chamber, a process called “freeze-etching,” the membrane surfaces come into view. If, following sublimation of the ice, the sample is now mechanically fractured by the impact of a microtome knife, a process called “freeze fracture” inner membrane surfaces and the fractured internal faces are exposed. The fracture generally takes place along a specific path, namely, the interface between the lipid leaflets that is shown by the thick dashed line in Fig. 18 (B), top, as the hydrophobic interior along this line has the least resistance to fracture. The fracture faces along the thick dashed line are now exposed, showing in the final electron micrograph a complex array of particles of different sizes and shapes as sketched in Fig. 18 (B), bottom. These intra-membrane particles have been found to be photosynthetically functional protein complexes residing in the thylakoid membrane. The two complementary fracture faces are called the E-face (EF) and P-face (PF), where “E” and “F” stand for exoplasmic and protoplasmic, respectively. The exoplasmic faces are in regions confined to the inner half, whereas the protoplasmic faces are in regions confined to the outer half of the thylakoid membrane. All thylakoid membranes have a protoplasmic leaflet (P) and an exoplasmic leaflet (E). Thus each leaflet has a true surface (S) and a fractured face (F). This terminology leads to the designations PS or ES for the true membrane surfaces and PF or EF for the fractured faces. In addition, depending on whether the thylakoid membrane comes from the stacked or unstacked regions, a subscript s or U is added to the designations for the surface and fractured face. When a replica of the fracture faces is viewed in the transmission electron microscope, such as the electron micrograph made for the barley thylakoid membrane at 100,000X magnification and shown in Fig. 18 (C), the four faces designated as and may appear to be side by side on the same plane in the figure, but actually the fracture path jumps from the middle of one membrane to the middle of an adjacent stacked membrane, as seen in Figs. 18 (B). In reality, the E- and P-faces are separated by a step equal to the thickness of the two leaflets, as indicated in the schematic drawing in Fig. 18 (B). The four fracture faces in Fig. 18 (C) show imbedded particles of various sizes. In particular, the face shows a profusion of ~16 nm-diameter particles. The complementary fracture face contains relatively tightly-packed, but rather indistinct and smaller particles. It is known that fractionation of a thylakoid membrane disrupted by detergent or by mechanical means yields separate granal and stromal fractions. The granal fraction has been found to be enriched in photosystem-II components and activity,

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while the stromal fraction is enriched in photosystem I. Extensive studies of the particles on the various fractured faces, correlated with biochemical studies of fractionated membrane components, showed that both the 16 nm-diameter particle and photosystem-II activity are located in the stacked lamellar region. In fact, it was found that in mutant plants and algae lacking Chl b, which is found principally in PS II, both LHC II and the large particles are missing in the face. These results led to the identification of the 16 nm-diameter particle as the PS-II reaction-center complex bound to a full complement of the light-harvesting complexes. Fig. 18 (D) summarizes the distribution of various particles in the fracture faces with known thylakoid protein complexes. Besides the association of the PS II/LHC II complex with the EF faces, was found to be distributed in both PF and EF faces, while the PS-I and the ATP-synthase complexes were associated with PF faces. We now examine some additional results also obtained by freeze-fracture electron microscopy using thylakoid membrane fragments. The membrane fragments were obtained either by detergent fragmentation and subsequent separation by differential centrifugation or by mechanical fragmentation followed by separation by “aqueous-polymer two-phase partition.” These studies yielded useful and complementary information on the thylakoid-membrane structure itself as well as the functional properties associated with the particular structure. Now we examine results from experiments by Dunahay, Staehelin, Seibert, Ogilvie and Berg38 using the oxygen-evolving (OE) chloroplast fragments prepared by the method of Berthold, Babcock and Yocum39 [the so-called “BBY” thylakoid or particle] and examined by freeze-fracture electron microscopy. Purified broken spinach chloroplasts for obtaining the BBY OE/PS-II thylakoids by washing and suspending them in a buffer containing 5 mM then treating them at a Triton Xl00-to-Chl ratio of 25 at 4 °C in darkness As depicted in the left part of Fig. 19 (A), detergent fragmentation of the unstacked granal end regions apparently leads to the formation of granal membrane fractions composed of pairs of appressed membrane fragments in which the lumenal surfaces and consequently the oxygen-evolving complexes exposed to the external environment (A) and (B) below]. The fact that the OE/PS-II preparation can be pelleted at relatively low centrifugation speed is consistent with the notion that the preparation represents highly enriched granal thylakoid fractions. It is important to note that the presence of 5 mM in the suspending medium is required to preserve the spatial differentiation of the thylakoid membranes, as a lower salt concentration leads to unstacking of the membranes and formation of particles of intermixed membranes. Freeze-fracture electron micrographs in Fig. 19 (B), top, show the freshly prepared BBY thylakoids by Triton treatment to be in the form of flat, double-membrane sheets with flat ends, resembling stacked granal membranes but without the margins of the original stack(ing), as depicted schematically in Fig. 19 (A), step When viewed toward the membrane face, typical and faces appear, but virtually no or faces [Fig. 19 (B) bottom]. The particles in the and faces have the same size and density as those of the unfractionated, control thylakoids, supporting the notion that the large particles of the PS-II/LHC-II complexes originated in the grana. Next we examine a chloroplast fragment with a unique architecture whose preparation was pioneered by Bertil Andersson and Hans-Erik Åkerlund and colleagues 40,41 . The fractionation procedure is briefly as follows. The chloroplasts are first fragmented by a shear force generated by extrusion through a needle valve in an apparatus called the “Yeda” press (“Yeda” is the name of the Israeli manufacturer) or the “French” press (named after Stacy French) [step in Fig. 19 (C)]. Again the chloroplasts must be placed in a medium containing simple cations, e.g., 5 mM or 150 mM NaCl, in order to preserve the pairing ofthe appressed membrane after mechanical disruption and fragmentation, so that the broken edges of the paired thylakoid membranes may then reseal to form an inside-out vesicle. Likewise, the stromal thylakoid membrane fragments also reseal but to form a right side-out vesicle in Fig. 19

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(C); actual forms shown after step The top box in Fig. 19 (C) shows the various structural features of the thylakoid membrane and the symbols and terms for various steps in the resealing processes. The resealed vesicles were then fractionated. Instead of using centrifugation, which is the most common method of fractionation and is based on the sizes and densities of the membrane fragments, the aqueous-polymer two-phase partition developed by Per-Åke Albertsson42 was used. This method separates the fragments on the basis of their surface properties in Fig. 19 (C)]. The partition system consists of two water-soluble polymers (dextran and polyethylene glycol) in a phosphate buffer that also contains various salts and either sucrose or sorbitol. A mixture of appropriate overall composition gives rise to two immiscible phases, each enriched in one or the other polymer and each phase being ~85-99% water. Attaining optimal phase composition requires consideration of the type and concentration of the polymers, ionic composition, temperature, etc. The phase-partition technique has since been successfully applied to the separation of various macromolecules, cells and organelles and their fragments and depends mainly on differences in surface properties of the constituents that are to be separated. Separation of the disrupted and resealed thylakoid-membrane vesicles is depicted in Fig. 19 (C), step About 2/3 of the thylakoid material is partitioned to the top phase T1 and 1/3 to the bottom phase B1. Each phase may be re-partitioned to further increasingly enrich one or the other type of vesicle, the number of re-partitions being designated by a number after T(op) or B(ottom). Typically, 55 and 20% of the thylakoid material is recovered in phases T2 and B3, respectively, as shown in Figure 19 (C). Åkerlund, Andersson and Albertsson43 analyzed the chemical composition and photochemical activity of the partitioned vesicles and found that the B3 vesicles were highly enriched in photosystem II, while the T2 vesicles were slightly enriched in photosystem I. Furthermore, the authors observed strikingly different behavior between the vesicles from the two phases with respect to light-induced proton translocation. As measured by a pH-electrode, the medium containing T2 vesicles showed a reversible lightinduced pH increase (loss of protons) in the medium, while the B3 vesicles showed a reversible pH decrease, or proton extrusion (gain in protons in the external medium), as shown in Figure 19 (C), right, immediately to the right of the sketch for each appropriate phase. Independent confirmation of the inside-out nature of the B3 vesicles was reported by Gräber, Zickler andÅkerlund 44 , who found that lightinduced charge separation across the membrane bilayer was exactly opposite to that found in normal thylakoids. Similar behavior in proton movement was also observed by measuring absorption changes of suitable pH-indicator dyes. A more direct confirmation of the inside-out character of the B3 vesicles was provided by freezefracture electron microscopy. Representative replicas ofthe T2 and B3 vesicles are shown in Fig. 19 (D). As expected for the B3 inside-out vesicles, large distinct particles appear in the EF faces next to the intra-thylakoid space [Fig. 19 (D), left], while closely packed small particles appear in the PF faces next to the stroma [Fig. 19 (D), right]. A large number of vesicular faces were examined and the results revealed that on average the B3 fraction contained 74% inside-out and 26% rightside-out vesicles while the T2 fraction contained 89% rightside-out and only 11% inside-out vesicles.

VIII. Composition of Thylakoid Membranes – Lipids, Proteins and Electron Carriers In this section we will discuss the three major classes of the various constituents of the thylakoid membrane, namely lipids, proteins and electron carriers. All the essential constituents are listed in Tables 1 to 5. Since most of these constituents or complexes will be discussed in more detail in later chapters, the tables simply serve as a convenient listing of these components for easy reference. A new volume by Siegenthaler and Murata45 should be consulted for the structure and function of lipids.

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The Lipids: Lipids, water-insoluble organic compounds, make up about 50% of the thylakoid mass. Thylakoid-membrane lipids have distinctly hydrophilic heads. The neutral galactolipids, monogalactosyl diacylglycerol (MGDG) and digalactosyl diacylglycerol (DGDG) together account for ~80% of the lipid composition of the thylakoid membrane, with the anionic sulfolipid, sulfoquinovosyl diacylglycerol (SQDG), and the phospholipid, phosphatidyl glycerol (PG), accounting for ~10% each. There may also be a small percentage of phosphatidyl choline, but its authenticity remains uncertain. The double envelope membranes of chloroplasts, on the other hand, are composed mostly of phospholipids. The four major thylakoid lipids, including their full names, abbreviations and structures are listed in Table 1. The galactolipids are known to be distributed asymmetrically in the lipid bilayer – about 60% of the galactolipids are present in the outer leaflets and 40% in the inner leaflets of the thylakoid-membrane bilayer. The strong interaction between the head-groups of the galactolipid molecules determine their packing properties and enhance the stability of the membrane. Formation of the bilayer structure of the thylakoid membrane also depends on the presence of proteins. Another characteristic of the thylakoid lipids is their high content of the trienoic acid, (C18:3) which contributes to the fluidity of the membrane, necessary for the diffusion of lipophilic compounds such as plastoquinone, and the lateral diffusion of protein complexes.

Protein complexes: The presence of protein complexes in the thylakoid membrane has been deduced from freeze-fracture electron microscopy as well as from biochemical studies. Each of the five major complexes contains multiple proteins and prosthetic groups. The number of proteins range from three in the LHC II complex to five or more complexes in cytochrome nine in the ATP-synthase complex and a little over 20 each in the PS-II and PS-I complexes. Many of the protein components will be discussed in detail in later chapters. For the present, we merely list the currently known protein components of the five complexes in Tables 2 to 5. The common names, molecular mass in kilodaltons, encoding genes, and some remarks about their functions or factors are also included in the tables. In addition,

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later chapters that deal specifically with the particular protein complex are indicated in the right margin. Most proteins in the PS-II complex are membrane spanning, but the three extrinsic proteins that are involved in oxygen evolution are located on the lumenal side of the thylakoid membrane. Although most proteins in the PS-I complex are also membrane-spanning, a few are located toward the stromal side of the thylakoid membrane, e.g., the iron-sulfur proteins that contain FeS-A and FeS-B and the Fd-docking protein. On the other hand, some mobile electron-transfer proteins are present on one side or the other of the membrane, with Fd present on the stromal side and near the ferredoxin(FNR) and the FNR-binding protein, and the copper-protein electron carrier, plastocyanin (PCy), present on the lumenal side, close to the PS-I primary donor P700. Note that LHCII is a separate light-harvesting complex, which supplements the inner antennae (CP29, CP26, CP24 and CP22) associated with the PS-II complex. The presence of this separate LHC II complex has been confirmed by the results of freeze-fracture experiments obtained with thylakoid membranes of both wild-type and mutant plants. The PS-II inner antenna complex and the LHC-II complex appear as distinctly different classes of membrane particles. On the other hand, the LHC-I proteins associated with the PS-I complex appear to be complexed to the PS-I reaction center, much as the inner antennae of PS II are complexed to the PS-II reaction center. Electron carriers and electron-transfer proteins: Electron-transfer reactions in photosynthesis involve electron carriers or electron-transfer proteins, including, among others, quinones, cytochromes, and ironsulfur proteins. In the following, we present a summary of the carriers or associated proteins that are primarily involved in photosynthetic electron-transfer reactions, along with a listing in Fig. 20. Although ATP is not an electron carrier, it is included in the figure to remind us of the common components present in the structures of ATP and molecules [see Fig. 20 (A)]. Photosynthetically active quinones include plastoquinone of green-plant photosystem II, ubiquinone and menaquinone in photosynthetic bacteria, and phylloquinone in photosystem I. Plastoquinone is present in green-plant photosystem II both as a tightly-bound and a loosely-bound electron carrier, designated and respectively. is photoreduced only to the semiquinone but can accept two electrons, forming the plastohydroquinone [see Chapters 5, 6 and 16 for further discussion]. Plastohydroquinone is the final reduction product of photosystem II and goes on to reduce the cytochrome complex as part of the electron transport and proton translocation processes [see Chapter 35 for detailed discussions]. Ubiquinone (UQ), also known as coenzyme Q, has a benzoquinone structure with a long side chain. The name “ubiquinone” is for the ubiquitous nature of the quinone. Some bacteria also contain menaquinone (vitamin either in addition to ubiquinone or in place of it. For instance, the reaction center of Rhodopseudomonas viridis contains one ubiquinone and one menaquinone, while in some other bacterial reaction centers both quinones are ubiquinones. Menaquinone has a naphthoquinone structure with a long isoprenoid side chain. The long hydrocarbon side chains in ubiquinone and menaquinone render a high degree of hydrophobicity to these molecules. All the quinones can undergo one-electron reduction to form a semiquinone which depending on the pH and the nature of the binding site, the semiquinone can either remain as an anion or become a neutral semiquinone, i.e., or Various possible pathways for reduction are illustrated with benzoquinone in Fig. 20 (B), right. The fully reduced hydroquinone may be non-protonated (di-anionic), singly protonated or fully protonated and electrically neutral. In photosystem I, the primary electron acceptor is a monomeric chlorophyll called The secondary electron acceptor is a bound phylloquinone (abbreviated as in this book), whose in situ quinone/ semiquinone redox potential has been estimated to be Phylloquinone has the same phytol side

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chain as chlorophyll. The phylloquinone acceptor will be discussed in detail in Chapter 33. Reduced phyllosemiquinone transfers an electron to an iron-sulfur protein called FeS-X, which has a redox-potential value of ~ -0.73 V. Redox-active amino acids are now recognized to play important roles in many biological electrontransfer reactions. In 1988, Bridgette Barry and Gerry Babcock46 used EPR spectroscopy to demonstrate the involvement of an isotope-labeled tyrosine radical in the water-splitting reaction in photosystem II of the cyanobacterium Synechocystis 6803. The manganese cluster directly responsible for the release of oxygen from water is apparently oxidized by via this redox-active amino acid [see Fig. 20 (C) left]. The oxidized tyrosine cation radical formed by is responsible for the “SII-type” EPR signal (see Chapter 22 for detailed discussion of this subject). The tyrosine cation radical has also been identified by optical spectroscopy47 as the immediate donor to (see Chapter 23). Tyr-161 of the Dl protein has been identified as the redox-active amino acid on the donor side of photosystem II by sitedirected mutagenesis (see Chapter 22, Section II.B. for details). As indicated above, tyrosine is able to transfer an electron to photooxidized in PS II. The oxidized tyrosine in turn can receive an electron coming from the splitting of water via the manganese cluster. The structure of the manganese cluster has been intensively studied by many including Ken Sauer and Mel Klein48 and their colleagues, using x-ray absorption edge spectroscopy (XAES) and extended x-ray absorption fine structure spectroscopy (EXAFS) [(see Chapter 20 for detailed discussion on the structure and function of the manganese cluster]. During the oxidation of water, the Mn-cluster is transformed step-wise from the initial oxidation state to four, increasingly higher oxidation S-states called the and the latter spontaneously reverting to the with the release of a molecule of oxygen. The change of the Mn oxidation states during S-state transitions will be discussed in Chapters 19 and 22.

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Cytochromes are versatile, electron-transfer proteins containing a heme, i.e., a macrocyclic tetrapyrrole coordinated to an iron atom, as the prosthetic group, similar to that found in hemoglobin and myoglobin. According to their characteristic absorption spectra and chemical composition, cytochromes may be divided into three groups, namely, cytochromes-a, -b and -c. Cytochromes transfer electrons with a change in the oxidation state between Fe (III) and Fe (II) in the heme. In photosynthetic organisms, mainly b- and c-type cytochromes are found. Although protoporphrin IX is the prosthetic group in both types [see Fig. 20 (D), left], there are slight differences in its composition. In the b-type cytochromes, both and are vinyl groups. The central iron atom, in addition to being coordinated by four N-atoms of the tetrapyrrole, is also axially ligated to two histidine residues from the protein [see Fig. 20 (D) center]. Cytochrome in particular, consists of two b-hemes, which allow the embedded cytochrome to provide a redox chain spanning the membrane. In a c-type cytochrome such as cytochrome [see Fig. 20 (D) right], the group of a protein cysteine residue is added to each of the two vinyl groups of the heme, thus covalently binding the heme to the protein through a sulfur bridge. In addition, the central Fe atom is axially coordinated to a histidine residue and to a terminal amino group of the protein [see Fig. 20 (D) right for cytochrome Iron-sulfur proteins, also called nonheme-iron proteins, are widely occurring, redox-active proteins, containing iron-sulfur clusters as the prosthetic groups. The clusters consist of equal numbers of iron and sulfide ions, with the iron atoms being coordinated to the protein through cysteine residues. The two most common types of iron-sulfur clusters, the [2Fe-2S] and [4Fe-4S] types, are each coordinated to four cysteine residues, and thus may be written as and respectively, as illustrated in Fig. 20 (E). Chloroplast ferredoxin containing the cluster is one common type of iron-sulfur protein. Another [2Fe-2S]-type protein is the Rieske iron-sulfur protein, present in the as well as the complex. The pair of iron atoms in the cluster of the Rieske iron-sulfur protein are bound to two cysteine and two histidine residues, in addition to two sulfur atoms. The three-dimensional structures of ferredoxins and that of the Rieske iron-sulfur protein have been determined by X-ray crystallography (see Chapters 34 and 35, respectively, for the structure of the chloroplast ferredoxin and the Rieske iron-sulfur protein). The sulfide ions in iron-sulfur proteins are acid-labile; this provides a simple means for detecting the iron-sulfur proteins, as the sulfide is released as upon acidification. The oxidized and reduced states of iron-sulfur clusters differ byjust one unit of formal charge, corresponding to and Iron-sulfurproteins are commonly characterized by optical absorption, circular-dichroism, EPR, Mössbauer spectroscopy, EXAFS etc. Plastocyanin is a low molecular-mass (10.5 kDa), copper-containing protein discovered by Sakae Katoh49 in Chlorella ellipsoidea in 1962. Plastocyanin is present in chloroplasts, as well as in cyanobacteria and red algae, but not in photosynthetic bacteria. When copper is lacking in the culture medium for some algae and cyanobacteria, for example, Scenedesmus, a soluble c-type cytochrome, which is similar to plastocyanin in molecular size as well as redox potential, may be formed as a substitute for plastocyanin. Its role is to mediate electron flow from cytochrome f of the complex, to of photosystem I. Electron transfer via plastocyanin occurs as a one-electron change in the oxidation state of its copper atom, which is coordinated at the active site by two nitrogen atoms from a pair of histidines and two sulfur atoms, one from a cysteine residue and the other from a methionine residue of the protein, as shown in Fig. 20 (F). Plastocyanin is a water-soluble protein, located in the lumen region and complexed to P700 through electrostatic attraction to facilitate rapid electron transfer to Plastocyanin will be presented in Chapter 34.

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IX. Lateral and Transverse Asymmetry of Thylakoid Membranes Freeze-fracture electron microscopy of thylakoid membranes has clearly revealed an asymmetric lateral distribution of the various photosynthetic complexes in the granal and stromal membranes, i.e., the distribution of the protein complexes in the membrane is nonrandom. This “lateral asymmetry” was further substantiated by the results of electron microscopy of the inside-out vesicles discussed in Section VII. These findings by electron microscopy are summarized by the model shown in Fig. 21 (A). It is a transverse cross section of the thylakoids shown earlier in Fig. 13 (D) and (D'), with the various photosynthetic protein complexes appropriately placed in the granal and stromal regions. As shown in both Fig. 21 (A) and (B), there are five major protein complexes in the thylakoid-membrane network: (1) the photosystem-II core with bound inner antennae (collectively designated as “PS II”), (2) the photosystem-I core with bound antennae LHC I (collectively designated as “PS I”), (3) the cytochrome complex, (4) the ATP-synthase and, finally (5) a separate, peripheral lightharvesting, chlorophyll-protein complex called “LHC II” that supplements the inner antennae bound to the photosystem-II reaction center . The model in Fig. 21 (A) shows that the PS-II complexes are present mainly in the appressed region of the thylakoid membrane, while the PS-I complexes are in the non-appressed, stromal region of the membrane. The are, on the hand, distributed almost equally among the appressed and non-appressed membranes. The ATP synthase complex is present exclusively in the stromal part of the thylakoid membrane. More than twenty years ago Miller and Staehelin 50 found that the ATPsynthase part of protein complex was lollipop-shaped, with the knob protruding from the non-appressed thylakoid membrane into the stroma. A typical distribution of the five major complexes between the appressed and non-appressed regions is summarized in the accompanying table in Fig. 21 (A). Of the five protein complexes, four are involved in the photochemical and electron-transport reactions while the fifth, the complex, uses a photoinduced, proton gradient to synthesize ATP. With this model we may go through the sequence of events in the photochemical and electron transport reactions taking place in green-plant photosynthesis. To facilitate the discussion, one segment of the thylakoid membrane containing the five protein complexes is depicted in Fig. 21 (B) along with the Z-scheme in Fig. 21 (C) incorporating the major reaction components. Absorption of a photon by the LHC-II/PSII complex initiates a charge separation to form the oxidized and the reduced by itself is used as the symbol for a pheophytin molecule in this book], through the manganese cluster i.e., the oxygen-evolving complex, and tyrosine, splits water molecules by successively extracting four electrons from two molecules of water to release one molecule of oxygen, and four protons into the (inside) lumen space [see Fig. 21 (B)]. The extra electron on is very rapidly transferred to a bound (primary) plastoquinone, designated as which in turn passes this electron to a loosely bound (secondary) plastoquinone [see Fig. 21 (C)]. For every pair of water molecules decomposed, two fully reduced and protonated plastoquinol molecules are formed. When has acquired two electrons from in two successive photoacts and extracted two protons ultimately from the stroma phase, it becomes the fully reduced plastohydroquinone (or plastoquinol), The PS-II complex is also called the“water-plastoquinone oxido-reductase.” A detailed discussion of the electron transfer reactions involving and of PS II is presented in Chapter 16.

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The

complex lying between PS II and PS I in the electron-transport system resembles the complex of mitochondria and photosynthetic bacteria. These cytochrome complexes possess one Rieske iron-sulfur protein “R-FeS” (a [2Fe-2S] protein discovered by John Rieske) and a so-called subunit IV. The two b-hemes of and the subunit IV span the thylakoid membrane, while the R-FeS and are located near the lumen side. As previously noted, the placement of the b-hemes across the thylakoid membrane helps form a redox chain across the membrane. The function of the complex in green-plant thylakoids is to oxidize the plastohydroquinone formed by PS II and to transfer these electrons to plastocyanin. Accordingly, the complex has therefore also been called the “plastohydroquinone-plastocyanin-oxidoreductase.” When the plastohydroquinone becomes fully oxidized at the lumenal surface of the membrane, it loses two electrons and also releases two protons to the (inside) lumen phase. Thus accompanying the reduction of one plastoquinone molecule by PS II and its subsequent reoxidation, there is a net transfer of two protons from the (outside) stromal phase into the (inside) lumen phase. Thus with the splitting of two water molecules by PS II to form one oxygen molecule, four protons are translocated across the membrane. As mentioned above, oxidation of two water molecules also releases an additional four protons into the lumen space. Thus water splitting and plastoquinone reduction/re-oxidation result in the generation of eight protons and the creation of a proton gradient across the thylakoid membrane. As indicated, above, the two b-hemes of the complex provide a pair of reacting sites spanning the thylakoid membrane, one near the stromal side and the other near the lumenal side of the thylakoid membrane. The plastohydroquinone is first oxidized by the Rieske FeS to a semiquinone, which is then oxidized by cytochrome f, which then releases the electron to the copper protein plastocyanin. After loss of one electron by the plastohydroquinone, the resulting semiquinone loses an electron to the two b-hemes in series. The b-hemes operate in the so-called “Q-cycle,” similar to that in the mitochondrial or bacterial cytochrome complex, and provide a translocation of additional protons across the membrane into the lumenal space. Discussion of the cytochrome complex and the Q-cycle will be presented in Chapter 35. At the time PS II absorbs a photon, PS-I complex also absorbs a photon, resulting in a charge separation to form the oxidized primary electron donor and the reduced primary electron acceptor The oxidized primary donor is then reduced by an electron from reduced plastocyanin. The electron on the primary acceptor is rapidly transferred step-wise through a series of acceptors consisting of a phylloquinone three iron-sulfur clusters in FeS-X, FeS-A and FeS-B, and finally ferredoxin (Fd) which reduces to NADPH in a reaction catalyzed by -ferredoxin reductase (FNR). The PS-I complex is also called the “plastocyanin-ferredoxin oxido-reductase.” The PS-II and PS-I complexes are linked by two mobile electron carriers: plastoquinone and plastocyanin to make a chain for electron transfers through the two photosystems, resulting in oxygen evolution at the PS-II end and reduction at the PS-I end. Electron transfer from PS II, through the Cyt complex and then to PS I follows a Z-shaped pathway, as shown by the thin-lined letter “Z” in Fig. 21 (B), passing through three of the complexes. The Z-scheme [also see discussion in section VI above] is a convenient formalism for describing the photosynthetic electron pathway. In reality, the supramolecular complexes are segregated into different regions of the thylakoid membrane and the so-called noncyclic electron transfer from PS II to PS I in the Z-pathway actually requires the small, mobile electron carriers to act as shuttles between the appressed and non-appressed regions of the thylakoids. Electrons released from the splitting of water on the lumen side (i.e., the inside) of the thylakoid membrane are passed through the electron-transport chain involving plastoquinone, the cytochrome complex and plastocyanin, to photosystem I and then to ferredoxin and eventually The overall electron flow is linear and vectorial, from the oxygen-evolving apparatus on the inside toward

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on the outside. The consequence of this vectorial electron flow, or displacement of electric charges, is the creation of an electric potential across the membrane, with positive charges on the inside and negative on the outside. A proton gradient is also developed across the membrane to create an additional electrostatic potential across the membrane. First, protons formed by the oxidation of water are released to the inside, resulting in an acidification of the thylakoid lumen space. In addition, protons are extracted from the outside, leading to the formation of and are ultimately released on the lumenal side during the reoxidation of this The end result is establishment of a proton gradient across the thylakoid membrane, which provides potential energy for doing chemical work. A fourth complex, the ATP synthase, utilizes the proton gradient and the membrane potential created to convert ADP into ATP. The ATP-synthase complex, present exclusively in the nonappressed thylakoid membrane facing the stroma, can be readily seen in electron micrographs as lollipop-shaped knobs protruding into the stroma. As noted, accompanying electron transfer in the chloroplast thylakoid membrane, there is proton transport from the stroma into the intra-thylakoid space, or lumen. Since the intra-thylakoid space is narrow and the volume of the flattened thylakoid lumen is small, even a small influx of protons results in a large change in proton concentration, i.e., in pH, in the lumen. For instance, a 10,000-fold difference ratio in proton concentration across the membrane, e.g., a stromal pH of 8 and a lumenal pH of 4, can provide a powerful driving force for ATP synthesis. To utilize the proton gradient formed during electron transport, the headpiece of ATP synthase must be oriented toward the stroma for protons in the lumen space to flow through to to achieve ATP synthesis.

X. Evolution of Photosynthetic Reaction Centers We have seen the Z-scheme for the two photosystems in green-plant photosynthesis and the electron carriers in these photosystems. We have also described how the photosystems of green plants and photosynthetic bacteria all appear to function with basically the same sort of mechanisms of energy transfer, primary charge separation, electron transfer, charge stabilization, etc., yet the molecular constituents of the two reaction centers in green plants, in particular, are quite different from each other. Photosystem I contains iron-sulfur proteins as electron acceptors and may thus be called the “iron-sulfur (FeS) type” reaction center, while photosystem II contains pheophytin as the primary electron acceptor and quinones as the secondary acceptors and may thus be called the “pheophytin-quinone type.” These two types of reaction centers have also been called RCI and RCII types, respectively. Interestingly, reaction centers of apparently all photosynthetic organisms may be assigned to one or the other of these two types. For instance, the reaction center of both purple bacteria and the green filamentous bacteria, Chloroflexaceae, and green-plant PS II are of the On the other hand, the green sulfur bacteria, Chlorobiaceae, the Heliobacteria, and photosystem I all have the FeS-type reaction centers. The interrelationship of the various organisms with respect to the type of reaction center(s) is shown in Fig. 22 by their electron-transport diagrams. The Z-shaped electron-transport diagram for green plant photosynthesis is placed in the center of the figure, where it is flanked by diagrams for the organisms with reaction centers of the same type. The individual electron carriers of the four types of reaction centers are also listed in the table in Fig. 22 (B), together with their redox potentials and reaction times. As we shall see below, the distinction between the two types of reaction centers is not absolute. For instance, both types of reaction centers are associated with peripheral light-harvesting, pigment complexes.

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The resemblance between PS II and the reaction centers of purple bacteria and filamentous green bacteria has been established by extensive biochemical and spectroscopic studies. In addition to the analogous electron carriers (bacterio)pheophytin and quinones present in purple bacteria and PS II, their reaction-center-core protein subunits, the L and M subunits in purple bacteria and the Dl and D2 subunits in PS II, show significant amino-acid sequence homologies. The recognition of this homology was initially pointed out by Achim Trebst51 on the basis of aminoacid sequence and greatly facilitated by the determination of the three-dimensional structure of a purple bacterium by X-ray diffraction by Deisenhofer, Epp, Miki, Huber and Michel 52 in 1985, and noted particularly by Hartmut Michel and Hans Deisenhofer53. Many of these features will be discussed in more detail in subsequent chapters. On the other hand, distinctions do exist between the purple bacterial and PS-II reaction centers. PS II contains but no comparable carrier is present in purple bacteria. The most outstanding distinction, however, is the unique ability of PS II to oxidize water. Although one may consider the purple bacteria to have a comparable ability to oxidize the mechanisms of the two processes are entirely different. Nevertheless, many of the structural similarities do suggest that the purple bacterial and PS-II reaction centers are evolutionarily related. The green sulfur bacteria, Chlorobiaceae, and the recently discovered heliobacteria possess a FeS-type reaction center, or RCI, similar to that in photosystem I of cyanobacteria and green plants. The heterodimeric core of the FeS-type reaction center in PS I is composed of two polypeptides but of comparable molecular weight and with a high percentage of amino-acid homology. On the other hand, the electron carriers in the reaction centers of Chlorobiaceae and Heliobacteriaceae, unlike PS I, are bound to a homodimeric reaction center. Still, both types of reaction centers contain a similar, dimericBChl primary electron donor associated with a monomeric-BChl primary electron acceptor The green sulfur bacteria, heliobacteria and PS I also contain a chain of virtually identical secondary and tertiary acceptors, including the iron-sulfur proteins FeS-X, FeS-A and FeS-B, with similar associated proteins and molecular orientations. Furthermore, the secondary electron acceptor is a phylloquinone in PS I and a rather similar naphthoquinone in the green sulfur bacteria and heliobacteria. A notable distinction is that the secondary electron donor side of the green sulfur bacteria and heliobacteria resembles that of the purple bacteria (a cytochrome) rather than that of PS I (plastocyanin). The diverse, present-day photosynthetic organisms with common structural and functional features suggest that they are probably derived from a single photosynthetic ancestor. Equally significant is that the evolutionary process must have, in time, refined the structural parameters for greater efficiency of the organisms. In the past, systematic classification of microorganisms was mainly based on morphological and physiological criteria. With the recent advances in molecular genetics, the nature of the evolutionary trail of the photosynthetic reaction centers can be reconstructed by studying the comparative biochemistry, amino-acid sequence analysis, or some other aspects of the existing organisms. Woese54 has reconstructed an evolutionary tree of the photosynthetic reaction centers based on 16S rRNA analysis. We present here, in Fig. 23, an evolutionary tree of the photosynthetic reaction centers presented by Schubert, Klukas, Saenger, Witt, Fromme and Krauß 55 that is based on the structural information for the PS-I reaction center as determined by x-ray crystallography at 4-Å resolution. The authors pointed out that the N-terminal antenna-binding domain and the reaction-center domain at the C-terminal of the FeS-type PS-I core proteins PsaA and PsaB are analogous to those of -type reaction center of purple bacteria. The analogy extends to the electron-transfer cofactors and their arrangement, the secondary structural motifs of the RC cores, and a significant amino-acid sequence homology between the N-terminal domains of the core proteins of RCI and the corresponding features of RCII. These analogies help support the hypothesis that both RCI and RCII share a common evolutionary origin.

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As already discussed above and shown in Fig. 23, all bacterial photosystems are anoxygenic and belong to either the FeS-type (RCI) or the (RCII). While the anoxygenic bacteria possess just a single RC-complex, of either the RCI or RCII type, oxygenic cyanobacteria, eukaryotic algae and green plants possess both RCI and RCII linked in series, splitting water at one end and reducing at the other.

For further reading: R1. RK Clayton (1980) Photosynthesis. Physical Mechanism and Chemical Patterns. Cambridge University Press R2. B Andersson and J Barber (1994) Composition, organization, and dynamics of thylakoid membranes. In: EE Bittar and J Barber (eds) Advances in Molecular and Cell Biology, 10: 1-53. JAI Press R3. LA Staehelin and GWM van der Staay (1996) Structure, composition, functional organization and dynamic properties of thylakoid membranes. In: DR Ort and CF Yocum (eds) Oxygenic Photosynthesis: The Light Reactions, pp 11-30. Kluwer R4. H Huzisige and B Ke (1993) Dynamics of the history ofphotosynthesis research. Photosynthesis Res 38: 185209 R5. Govindjee (2000) Milestones in photosynthesis research. In: U Pathre, H Younis and PK Mohanty (eds) Probing Photosynthesis, pp 9-39. Taylor and Francis (UK)

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References: 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

15. 16.

17. 18. 19 . 20. 21.

22. 23. 24. 25. 26. 27. 28. 29. 30. 31.

VM Albers and HV Knorr (1937) Absorption spectra of single chloroplasts in living cells in the region from 664 nm to 704 nm. Plant Physiol 12: 833-843 CS French (1958) Various forms of chlorophyll a in plants. Brookhaven Symp Biol 11: 65-73 CS French (1957) Derivative spectrophotometry. In: Proc l.S.A. Instrument and Control Symp, pp 83-94. Northern California Section, Instrument Society of America, Berkeley, CA M Havaux (1998) Carotenoids as membrane stabilizers in chloroplasts. Trends in Plant Sci. 3: 147-151 R Willst tter and A Stoll (1918) Untersuchungen ber die Assimilation der Kohlens ure. Springer, Berlin O Warburg (1919) ber die Geschwindigkeit der photochemischen Kohlens urezersetzung in lebenden Zellen. Biochem Z 100: 230-270 CB van Niel (1929) Photosynthesis in bacteria. In: Contribution to Marine Biology, pp 161-169. Stanford University CB van Niel (1941) The bacterial photosynthesis and their importance for the general problem of photosynthesis. Adv Enzymology 1: 263-328 S Ruben, M Randall, MD Kamen and JL Hyde (1941) Heavy oxygen as tracer in the study ofphotosynthesis. J Am Chem Soc 63: 877-878 MD Kamen (1963) Primary Processes in Photosynthesis. Acad Press R Emerson and W Arnold (1932) A separation of the reactions in photosynthesis by means of intermittent light. J Gen Physiol 15: 391-420 R Emerson and W Arnold (1932) The photochemical reaction in photosynthesis. J Gen Physiol 16: 191-205 Govindjee (1999) On the requirement of minimum number of four versus eight quanta of light for the evolution of one molecule of oxygen in photosynthesis: A historical note. Photosynthesis Res 59: 249-254 RE Fenna, BW Mathews, JM Olson and EK Shaw (1974) Structure of a bacteriochlorophyll-protein from the green photosynthetic bacterium Chlorobium limicola: crystallographic evidence for a trimer. J Mol Biol 84: 231240 DE Tronrud, MF Schmid and BW Mathews (1986) Structure and x-ray amino acid sequence of a bacteriochlorophyll a protein from Prosthecochloris aestuarii refined at 1.9 ¯ resolution. J Mol Biol 188: 443-454 T Katoh, M Mimuro and S Takaichi (1989) Light-harvesting particles isolated from a brown alga, Dictyota dichotoma. A supramolecular assembly of fucoxanthin-chlorophyll-protein complexes. Biochim Biophys Acta 976: 233-240 T Katoh, A Tanaka and M Mimuro (1993) Xanthosomes: supramolecular assemblies of xanthophyll-chlorophyll a/c protein complexes. Methods in Enzymology 214: 402-412 T Ogawa, F Obata and K Shibata (1966) Two pigment proteins in spinach chloroplasts. Biochim Biophys Acta 112: 223-234 R Hill (1939) Oxygen produced by isolated chloroplasts. Proc Roy Soc B127: 192-210 W Menke (1962) Structure and chemistry of plastids. Annu Rev Plant Physiology 13: 27-44 RB Park and NG Pon (1961) Correlation ofstructure with function in Spinacea oleracea chloroplasts. J Mol Biol 3: 1-10 R Emerson and CM Lewis (1943) The dependence of the quantum yield of Chlorella photosynthesis on wave length of light. Am J Bot 30: 165-178 T Tanada (1951) The photosynthetic efficiency of carotenoid pigments in Navicula minima. Am J Bot 38: 276283 FT Haxo and LR Blinks (1950) Photosynthetic action spectra of marine algae. J Gen Physiol 33: 389-422 R Emerson, R Chalmers and C Cederstrand (1957) Some factors influencing the long-wave limit of photosynthesis. Proc Nat Acad Sci, USA 43: 133-143 R Hill and F Bendall (1960) Function of two cytochrome components in chloroplasts: A working hypothesis. Nature 1186: 136-137 R Hill and R Scarisbrick (1951) The haematin compounds of leaves. New Phytol 50: 98-111 R Hill (1954) The cytochrome b component of chloroplasts. Nature 174: 501-503 R Hill (1965) The biochemist’s green mansions: the photosynthetic electron-transport chain in plants. In: PN Campbell and GD Greville (eds) Essays in Biochemistry 1: 121-151. Acad Press, London Govindjee and E Rabinowitch (1960) Two forms of chlorophyll a in vivo with distinct photochemical function. Science 132: 355-356 Govindjee, S Ichimura, C Cederstrand and E Rabinowitch (1960) Effect of combining far-red light with shorter wave light on the excitation of fluorescence in Chlorella. Arch Biochem Biophys 89: 322-323

46 32. H Kautsky, W Appel and H Amann (1960) Chlorophyllfluorescenz und Kohlens ureassimilation. XIII. Mittei-

lung. Die Fluorescenzkurve und die Photochemie der Pflanze. Biochem Z 332: 277-292 33. B Kok and G Hoch (1961) Spectral changes in photosynthesis. In: WD McElroy and B Glass (eds) Light and

Life, pp 397-416. Johns Hopkins Press 34. LNM Duysens, J Amesz and BM Kamp (1961) Two photochemical systems in photosynthesis. Nature 190:

510-511 35. HT Witt, A M ller and B Rumberg (1961) Experimental evidence for the mechanism of photosynthesis. Nature

191: 194-195 36. HT Witt, A M ller and B Rumberg (1961) Oxidized cytochrome and chlorophyll in photosynthesis. Nature 192:

967-969 37. B Rumberg, P Schmidt-Mende, U Siggel, B Skerra and HT Witt (1965) Analyse der Photosynthese mit Blitzlicht.

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IV. Quantitative Kopplung der Reaktionscyclen I und II im vollst ndigen Reaktionssystem. Z Naturforsch 20b: 1102-1116 TG Dunahay, LA Staehelin, M Seibert, PD Ogilvie and SP Berg (1984) Structural, biochemical and biophysical characterization of four oxygen-evolving photosystem II preparations from spinach. Biochim Biophys Acta 764: 179-193 DA Berthold, GT Babcock and CF Yocum (1981) A highly resolved oxygen-evolving photosystem II preparation from spinach thylakoid membranes. FEBS Lett 134: 231-234 B Andersson and H-E ¯kerlund (1978) Inside-out membrane vesicles isolated from spinach thylakoids. B i o chim Biophys Acta 503: 462-472 B Andersson and JM Anderson (1980) Lateral heterogeneity in the distribution of chlorophyll-protein complexes of the thylakoid membranes of spinach chloroplasts. Biochim Biophys Acta 593: 427-440 P-¯ Albertsson (1985) Partition of Cell Particles and Macromolecules (3rd edition) John Wiley H-E ¯kerlund, B Andersson and P-¯ Albertsson (1976) Isolation of photosystem II enriched membrane vesicles from spinach chloroplasts by phase partition. Biochim Biophys Acta 449: 525-535 P Gr ber, A Zickler and H-E ¯kerlund (1978) Electric evidence for the isolation of inside-out vesicles from spinach chloroplasts. FEBS Lett 96: 233-237 P-A Siegenthaler and N Murata (199 ) Lipids in Photosynthesis: Structure, Function and Genetics. Kluwer BA Barry and GT Babcock (1987) Tyrosine radicals are involved in the photosynthetic oxygen-evolving s y stem. Proc Nat Acad Sci, USA 84: 7099-7103 S Gerken, K Brettel, E Schlodder and HT Witt (1988) Optical characterization of the immediate donor to chlo rophyll in -evolving photosystem-ll complexes. FEBS Lett 237: 69-75 K Sauer, VK Yachandra, RD Britt and MP Klein (1992) The photosynthetic water oxidation complex studied by EPR and x-ray absorption spectroscopy. In: VL Pecararo (ed) Manganese Redox Enzymes, pp 141-175. VCH Publ S Katoh (1960) A new copper protein from Chlorella ellipsoidea. Nature 186: 533-534 KR Miller and LA Staehelin (1976) Analysis of the thylakoid outer surface: Coupling factor is limited to unstacked membrane regions. J Cell Biol 68: 30-47 A Trebst (1986) The topology of the plastoquinone and herbicide binding peptides of photosystem II in the thylakoid membrane. Z Naturforsch 41c: 240-245 J Deisenhofer, O Epp, K Miki, R Huber and H Michel (1984) X-ray structure analysis of a membrane protein complex. Electron density map at 3 ¯ resolution and a model of the chromophores of the photosynthetic reaction center from Rhodopseudomonas viridis. J Mol Biol 180: 385-398 H Michel and J Deisenhofer (1988) Relevance of the photosynthetic reaction center from purple bacteria to the structure of photosystem II. Biochemistry 27: 1-7 CR Woese (1987) Bacterial evolution. Microbiol Rev 51: 221-271 W-D Schubert, O Klukas, W Saenger, HT Witt, P Fromme and N Krau (1998) A common ancestor for oxygenic and anoxygenic photosynthetic systems: A comparison based on the structural model of photosystem I. J Mol Biol 280: 297-314

Chapter 2

The Bacterial Photosynthetic Reaction Center: Chemical Composition and Crystal Structure Structure of the Reaction Centers of Photosynthetic Bacteria A. Protein Structure B. Amino-Acid Sequences of Rb. sphaeroides and Rp. viridis Reaction-Center Polypeptides C. Hydropathy Plots for the Bacterial Reaction-Center Polypeptides II. Three-Dimensional Structure of Bacterial Photosynthetic Reaction Centers A. Crystallization B. Crystal Structure of Protein Subunits, Pigment Molecules and Electron Carriers 1. Rp. viridis 2. Rb. sphaeroides 3. Spatial Arrangement of the Polypeptide Subunits Summary Reaction-Center Structure and Photochemical Reactions III.

l.

48 49 50 51 54 54 56 56 58 59 62

Photosynthetic prokaryotes such as cyanobacteria and photosynthetic bacteria lack chloroplasts and in these organisms the light reactions that drive photosynthesis take place in the cell’s inner plasma membrane. The photosynthetic apparatus of purple bacteria, for example, is contained in a system of “intracytoplasmic membranes.” Fig. 1 depicts the morphologies of two such purple bacteria – Rhodobacter (Rb.) sphaeroides [Fig. l (A)], formerly called Rhodopseudomonas sphaeroides, and Rhodopseudomonas (Rp.) viridis [Fig. 1 (B)] – species that are commonly used for photosynthesis studies. The former contains bacteriochlorophyll a (BChl a), which absorbs in the 800-880 nm region in vivo, while the latter contains BChl b, which absorbs in the 960-1020 nm region. Figs. 1 (A) and (C) show how the parent cytoplasmic membrane in Rb. sphaeroides has differentiated into the specialized intracytoplasmic membrane by repeated invagination, as pointed to by a small arrow at the cell wall. During the process of invagination, the region outside the cytoplasmic membrane, i.e., the periplasm between the cytoplasmic membrane and the cell wall, becomes virtually trapped inside the newly formed cavity. When bacterial cells are broken, for example by disruption through a French pressure cell or by sonication, both the cytoplasmic and intracytoplasmic membranes are fragmented but the fragments reseal to form quasi-spherical vesicles called “chromatophores.” In Rp. viridis cells, on the other hand, the cytoplasmic membrane invaginates by folding back and forth to form stacked lamellae [Fig. 1 (B)]. In some species of photosynthetic bacteria, the invaginated cytoplasmic membranes display still other morphologies. Fig. 1 (C) is a highly-idealized sketch showing the reaction-center core with light-harvesting complexes and the ATPase embedded in the cytoplasmic membrane and the orientation of each with respect to light-promoted proton flux. Fig. 1 (D) shows a detailed model of the photosynthetic membrane of Rb. capsulatus with its various components embedded in the phospholipid matrix of the intracytoplasmic membrane.

48

I. Structure of the Reaction Centers of Photosynthetic Bacteria The first photosynthetic reaction center to be isolated was that reported by Reed and Clayton1 for the mutant strain R-26 of Rb. sphaeroides by Triton treatment. Other methods for isolating reaction centers were subsequently developed by Clayton and Wang2, Feher and Okamura 3 , Gingras4 and by others. Currently, pure reaction centers can be isolated from most photosynthetic bacteria, including Chlorobium as reported recently by Hager-Braun et al5. The basic procedure for isolating reaction centers from the bacterial cells is to first break the cells and separate the chromatophore components by differential centrifugation. The membrane components may then be purified further either by density-gradient centrifugation, precipitation with ammonium sulfate, DEAE (diethylaminoethyl cellulose) chromatography, gel filtration, dialysis, etc. It is worth noting that most reaction-center preparations still retain photochemical activity. For instance, cytochromes present in the isolated reaction-center preparation from the purple bacterium Chromatium vinosum can still supply electrons to the photooxidized primary donor, even at cryogenic temperatures. We will now discuss the chemical composition and the threedimensional structure of reaction centers of two photosynthetic purple bacteria, namely, Rp. viridis and the carotenoidless mutant of Rb, sphaeroides R-26.

Chapter 2

The Bacterial Photosynthetic Reaction Center

49

I. A. Protein Structure The chemical composition of the highly purified reaction-center complex from several photosynthetic bacteria is now well established. In addition to protein subunits designated as “L” (light), “M” (medium) and “H” (heavy), some reaction centers also contain a cytochrome (C) subunit. The chemical composition of reaction-center complexes of several of these bacteria is shown in Table 1:

The purified reaction-center complex of Rb. sphaeroides R-26 can be separated by SDS-PAGE (sodium dodecyl sulfate polyacrylamide-gel electrophoresis) into three polypeptides [Fig. 2 (A)]. The three fractions were named the L-, M- and H-subunits, representing light, medium and heavy, respectively, according to the migration distance of each in the electrophoresis gel. The molecular masses of the three fractions were originally estimated to be ~21 kDa, ~24 kDa, and ~28 kDa for the L, M and Hsubunits, respectively. Later the molecular-weight values estimated from gel migration were found to be incorrect because, apparently, the SDS-PAGE technique is sensitive to the hydrophobicity of the proteins being analyzed. The value of 28-kDa estimated for the more hydrophilic H-subunit turned out to be nearly correct but those original estimates of the more hydrophobic L- and M-subunits were ~30% lower than their actual values, based on the amino-acid composition derived from gene sequences. The correct molecular-weight values for the L-, M-, and H-subunits are actually 31, 34 and 28 kDa, as will be discussed below. Although no longer appropriate, the original designation L-, M- and H-, for each subunit, however, is retained. The three protein subunits are present at a 1:1:1 stoichiometry in the reaction center. The H-subunit can be readily removed from the reaction-center complex by washing with any of a variety of detergent solutions. For example, when the reaction-center complex is treated with a solution containing 1% SDS and 0.6% LDAO (lauryl N,N-dimethylamine-N-oxide), it is separated into the Hsubunit and an LM-complex. Figs. 2 (B) and (C) show the SDS-PAGE patterns obtained with the LMcomplex and the H-subunit subfraction, respectively. It is seen that the LM-complex can be further resolved into the L- and M-subunits and that all three subunits migrate to their proper locations. This LM-complex prepared with LDAO was found to be photochemically active at ambient temperature but not at low temperatures, but an LM-complex prepared later with the chaotropic agent was found to be photochemically active at both ambient and low temperature. The LM-complex contains one ubiquinone molecule and one Fe atom; it is associated with reduction of the quinone electron acceptor as evidenced by the EPR signal. These results suggest that the LM-complex may be the smallest photochemically active unit of the photosynthetic bacteria.

50

I. B. Amino-Acid Sequences in Rb. sphaeroides and Rp. viridis Reaction-Center Polypeptides The amino-acid composition of the L-, M- and H-subunits of Rb. sphaeroides and Rp. viridis has now been derived from the nucleotide sequence of the structural genes that encode the polypeptide subunits. The number of amino-acid residues and the precise molecular masses of the L-, M- and H-subunits in Rb. sphaeroides and Rp. viridis are listed in Table 2. Note that for verification purposes, selected polypeptides obtained by tryptic-digestive fragmentation are often subjected to amino-acid sequence analysis.

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As noted above, the molecular masses of the L- and M-subunits are actually higher than those originally estimated from SDS-PAGE and the H-subunit turns out to be the smallest among the three. The Land M-subunits are highly hydrophobic, each containing ~70% nonpolar amino acids, while the polar amino-acid content in the H-subunit is higher than in either the L- or M-subunit. The L- (and H-)subunit contains three cysteines, but the M-subunit contains none. It is possible to utilize this difference to separate the L- and M-subunits by means of affinity chromatography. Fig. 3 shows the amino-acid sequences of the three polypeptide subunits in reaction centers from the photosynthetic bacteria Rb. sphaeroides and Rp. viridis. Starting from the amino terminus of the L- and M-subunit sequences, every tenth amino-acid residue is marked by a dot above the sequence and numbered, the number here referring to Rb. sphaeroides. Two prominent features stand out in the amino-acid sequence. First, the amino-acid sequences of the L- and M-subunits from the two species are highly conserved, as are the H-subunits, as can be seen in the match-up ofthe residues shown underlined in Fig. 3 (A). By introducing gaps in the sequences, the alignment of the amino-acid sequences can be optimized to show maximum homology. The L- and M-subunits of the two species are homologous to each other. The percentage of identical residues between the L-subunit of Rb. sphaeroides and Rp. viridis is 59% and for the M-subunits 50%. There is an even higher percentage identity in the L- and M-subunits among Rb. sphaeroides and Rps. capsulata. Inspection of the highly conserved regions also reveals that these regions contain a high proportion of hydrophobic residues that are critical for their structural roles. Five regions in these two polypeptides contain at least 19 hydrophobic amino acids each, thus allowing each of them to form transmembrane helices. As seen from the hydropathy plots in Fig. 4 below and from direct structure determinations, this is indeed the case. The H-subunits of Rb. sphaeroides and Rp. viridis have a 39% subunit identity, and each has a region with a sufficient number of hydrophobic residues to form one transmembrane helix. Further details will be discussed in the following sections on the hydropathy plots and crystal structure of the bacterial reaction centers. These homologies indicate an evolutionary connection among the subunit proteins of the various species. The homology between the L- and M-subunits among Rb. sphaeroides and Rp. viridis [as well as Rp. capsulata and Rs. rubrum (not shown)] suggests that the genes for the L- and M-subunits arose from duplication of a single ancestral gene.

I.C. Hydropathy Plots for the Bacterial Reaction-Center Polypeptides The configuration of a protein in the membrane depends on the degree of hydrophilicity versus hydrophobicity of the amino-acid residues along the polypeptide chain. In general, a hydrophilic part will be exposed to the aqueous phase, while a hydrophobic part will be found buried inside the membrane, possibly in close contact with the hydrocarbon chains of the phospholipid bilayer. The hydropathy plot, generated by a simple yet elegant program devised by Kyte and Doolittle6 in 1982, shows the relative hydrophilicity and hydrophobicity of the amino acids along a polypeptide sequence. The word “hydropathy” which can be taken to mean “strong feeling about water,” really refers to both hydrophilic and hydrophobic character. The program uses a hydropathy scale in which each amino acid is assigned a value reflecting its relative hydrophilicity or hydrophobicity. The scale ranges from 4.5 for isoleucine, judged to be the most hydrophobic amino acid, to -4.5 for arginine, the most hydrophilic residue. A moving “window” consisting of a specified number of amino-acid residues is stepped through the aminoacid sequence and the average hydropathy of the residues within the window is determined at each step. The consecutive scores are then plotted from the amino terminus to the carboxy terminus, with the grand average of the hydropathy of the amino-acid composition of the sequenced protein taken as the mid-

52

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53

point. Kyte and Doolittle demonstrated the validity of the method by comparing plots for proteins with known structures determined by x-ray crystallography, and found the procedure to be a valuable tool for distinguishing exterior from interior portions of a protein and for predicting its structure within a membrane. Figure 4 shows the hydropathy plots for the L-, M- and H-polypeptides of Rb. sphaeroides R-26, using a moving window of 19 amino-acid residues for each scan. The plots for the L- and M-polypeptides [Fig. 4 (A) and (B)] are very similar, showing that both polypeptides have five hydrophobic segments. Each of these hydrophobic segments contains enough amino acids to form a membrane-spanning Note that the residue-number scale strictly applies only to the L- and H-polypeptides; that for the M-polypeptide has been shifted in order to maximize the coincidence of its hydrophobic regions with those of the Lpolypeptide. The hydropathy plot for the H-polypeptide [Fig. 4 (C) ] shows that it has only one hydrophobic region, indicating the presence of just one Thus the presence of eleven transmembrane helices in the L-, M- and H-subunits as predicted by hydropathy plots is in accord with conclusions drawn from previous studies by circular dichroism and polarized infrared spectroscopy and later confirmed by X-ray diffraction studies.

54

II. Three-Dimensional Structure of Bacterial Photosynthetic Reaction Centers Since membrane proteins are usually embedded in a lipid bilayer, their isolation generally requires solubilization with detergents and as a group they have not been crystallized until very recently. The difficulty has been that detergent molecules were randomly attached to the isolated proteins, and thus identical units were not available for producing an ordered crystalline lattice. Although several hundred different proteins have now been crystallized and their crystal structures determined, only a few of them are membrane proteins. These include the reaction-center complexes from the BChl b-containing photosynthetic bacterium Rp. viridis and the BChl a-containing bacterium Rb. sphaeroides. In 1982, Hartmut Michel7 succeeded in obtaining crystals of the reaction center of Rp. viridis and, soon after, Deisenhofer, Epp, Miki, Huber and Michel 8 used X-ray diffraction to determine its threedimensional structure. These results have been recognized as one of the most spectacular and important developments in the history of photosynthesis research. Before satisfactory crystals became available and X-ray diffraction could be used to directly yield the precise information on the structure of the reaction centers, all information about the reaction center could be derived only by indirect means. For example, the presence of predicted by hydropathy plots could now be determined directly by X-ray diffraction. In the following, we will first describe how the crystals are prepared, and then show the results of the X-ray crystallographic determination.

II.A. Crystallization To prepare membrane-protein crystals one must have very pure starting materials and, fortunately, reaction-center complexes of high purity have been available since the early 1980s for many photosynthetic bacteria. The rationale for the techniques used in preparing crystals of membrane proteins will be described first. Membrane proteins can be solubilized into aqueous solution with a detergent, as shown in Fig. 5, but when the detergent is completely removed, the protein becomes aggregated as a result of hydrophobic interactions and does not form crystals. Detergent molecules, like lipids, are amphiphilic and, like lipids, above a certain concentration, the so-called “critical micellar concentration,” they aggregate to form micelles, in which the hydrocarbon chains of the detergent molecules are oriented toward the inside, keeping them from making contact with the water molecules. In fact, when proteins are solubilized by detergents, the lipid molecules are essentially replaced by detergent molecules, with the hydrocarbon chains of the detergent molecules coming into contact with the hydrophobic part of the protein to form a “compound” micelle (see Fig. 5, bottom).

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The protein partially surrounded by detergent molecules can be used as the starting material for crystallization. Bringing a protein-detergent micelle into a three-dimensional crystalline lattice requires consideration of its hydrophobic and hydrophilic properties. In principle, there are two possible ways of arranging membrane proteins into a three-dimensional crystal, as shown by the cross-sectional views of the so-called “type-I” and “type-II” crystals in Fig. 6. Fig. 6, left shows an end view of a type-I crystal formed by stacking two-dimensional “crystal layers,” ordered sheets of proteins. Many proteins, but not all, can form such a two-dimensional crystal layer, in which the hydrophobic regions of the proteins interact with the hydrocarbon tails of the lipids, the twodimensional structure being stabilized by both hydrophobic and polar interactions. In each two-dimensional crystal layer no detergent is present and only the polar domains are exposed at the surface. These two-dimensional crystal layers then stack up to form a three-dimensional crystal through polar attractions between the layers. In three-dimensional crystals, the successive two-dimensional crystal layers need to be ordered in the third dimension with respect to translation, rotation and up-down orientation. Examples of type-I crystals which have been prepared are mitochondrial cytochrome oxidase, chloroplast Chl-a/b proteins, and a protein from the purple membrane of halobacteria. Two-dimensional crystals are usually rather small and useful only for examination by electron microscopy.

The “type-II” crystal shown in Fig. 6, right represents an alternative way to form membrane-protein crystals by having the proteins partially surrounded by detergent molecules. Type-II crystals are primarily stabilized by attractive forces between the polar domains of the proteins. Clearly, in order to accommodate the detergent molecules and still have a large polar attraction between protein molecules, it would be easier for proteins with large polar domains to form this type of crystals than those with small polar domains. In any event, in forming a type-II crystal, the size of the detergent molecules and hence of the micelle is of critical importance. For instance, large detergent molecules might bulge out into the polar domains of the protein, thus diminishing the polar interactions necessary for crystal formation. Thus, micelles formed by small detergent molecules might be needed so that the hydrocarbon tails of the detergent molecules just fit into the available space in the hydrophobic cavities formed by the meeting of the polar domains of the protein molecules. However, in practice, proteins are less stable with detergents having rather short hydrocarbon chains. Consequently, a compromise has to be made in this regard. For example, for Rp. viridis reaction-center proteins, detergents such as LDAO turned out to be satisfactory. Michel introduced an ingenious and effective means for modifying and controlling the detergentmicelle size by adding small amphiphilic molecules, the most effective among them being heptane-

56

1,2,3-triol. This small amphiphilic molecule itself is not a detergent, as its hydrocarbon chain is too short to form micelles. However, it can form mixed micelles with the detergent molecules and the resulting micelle is smaller than those formed with detergent molecules alone. These small amphiphilic molecules may also replace certain detergent molecules which might disturb the crystalline order. For instance, some detergent molecules (such as octylglucoside with a large glucose head group) might be too large to permit the formation of a protein crystal lattice. The polar head of a small amphiphilic molecule however is smaller than that of a detergent molecule and thus less of the polar domain of the protein would be covered and consequently more of it available for crystal formation. That replacement of detergent molecules with small amphiphilic molecules is favorable can be argued on more specific energetic grounds in that the energy gained in the formation of additional polar bonds during crystallization is greater than that needed to replace the detergent molecules by small amphiphilic molecules. Michel succeeded in crystallizing type-II crystals from the reaction-center protein of Rp. viridis in 1982. Crystallization was achieved by vapor diffusion using a concentrated reaction-center solution (1.5 mg protein per mL) containing 1.5 M ammonium sulfate, 3% heptane-1,2,3-triol, 3% triethylammonium phosphate and 0.5% LDAO in osmotic equilibration vs. a large reservoir of 2-3 M ammonium sulfate solution to attain supersaturation of the protein solution. Initially crystals as large as 0.4 mm X 0.4 mm X 1 mm were obtained within three weeks at room temperature and proved suitable for high-resolution structure determination by X-ray crystallography. Notably, the reaction-center crystals still retained photochemical activity. Both the isolated and the crystallized reaction centers of Rp. viridis contain four protein subunits, including the membrane-bound cytochrome subunit. Crystallization of the reactioncenter proteins from Rb. sphaeroides was subsequently achieved by Allen and Feher9, by Chang, Schiffer, Tiede, Smith and Norris10 and by Arnoux, Gaucher, Ducruix and Reiss-Husson11 and studied by X-ray diffraction crystallography.

II.B. Crystal Structure of Protein Subunits, Pigment Molecules and Electron Carriers With sufficiently large crystals of the bacterial reaction centers available, the determination of crystal structure by X-ray diffraction became possible and in 1983, Deisenhofer, Epp, Miki, Huber and Michel 8 determined the crystal structure of the Rp. viridis reaction centers at 3 Å resolution [later refined to 2.3 Å12]. From the electron-density map, the spatial arrangement of the polypeptide subunits, the pigment molecules and the electron carriers in the reaction center was determined. For this work, Deisenhofer, Michel and Huber were awarded the Nobel Prize in 1988.

II.B.1. Rp. viridis As previously noted, the Rp. viridis reaction center consists of four polypeptides: the L-, M-, H- and Cpolypeptides (C for c-type cytochrome), as shown in the model in Fig. 7. Each of the L- and M-polypeptides contains five transmembrane helices, labeled A to E and single- and double-underlined. The single H-subunit helix is triple-underlined. As mentioned earlier, the number of transmembrane helices determined by X-ray crystallography agrees with that predicted from the hydropathy plots for all three subunits (see Fig. 4). The helical content in the LMH complex is 51%, in good agreement with the 50±10% value obtained from circular-dichroism measurements13. The ten helical segments from the L- and M-subunits, together with the one helical segment near the N-terminus of the H-polypeptide, form an elliptical hydrophobic cylinder 45 Å-long, with major and minor diameters of 70 and 30 Å, respectively. The cylinder is embedded in the lipid-bilayer membrane and forms a framework for organizing and positioning the cofactors. The four D- and E-segments of the

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L- and M-polypeptide subunits form the core structure and are closely associated with the cofactors, while the A-, B- and C-segments of both the L- and M-subunits are located on the periphery of the reaction center. Some of the polypeptide segments connecting the transmembrane helices in the L- and M-polypeptides contain short helices which lie approximately parallel to the membrane plane and form part of the flat surfaces at the top and the bottom of the cylinder. The C-terminal half of the H-polypeptide forms a globular domain that attaches to the LM core on the cytoplasmic side of the membrane. The cytochrome subunit located on the periplasmic side and the globular domain of the H-subunit are both polar. From the tip of the cytochrome subunit to the bottom ofthe globular domain of the H-subunit, the reaction center measures 130 Å. Each reaction center contains 14 cofactors (shown by black rectangles, diamonds, etc. in Fig. 7): four BChl b, two one carotenoid (not shown), two quinones, one nonheme iron atom lying between the two quinones, and four cytochrome-c heme groups in the C-subunit. Sometimes, only the menaquinone on the L-side is retained in the reaction-center crystal, the other quinone being lost during purification ofthe reaction-center proteins. The L- and M-subunits with their cofactors are nearly symmetrical with respect to the axis perpendicular to the membrane plane. The cytochromeand the H-subunits, on the other hand, do not obey the local symmetry of the central portion of the reaction center, although the cytochrome subunit does have internal local symmetry of its own.

58

The cytochrome subunit (see Fig. 7, top), the largest subunit in the RC complex of Rp. viridis, contains 336 amino-acid residues. The cytochrome subunit sits on a flat surface on the periplasmic side ofthe LM core and consists of two pairs of heme-binding segments. Each heme-binding segment consists of a helix with an average length of 17 amino-acid residues.

II.B.2. Rb. sphaeroides Using X-ray diffraction, George Feher and determined the crystal structure of the Rb. sphaeroides reaction-center complex soon after that of Rp. viridis had been reported. Although the reaction center of Rb. sphaeroides had already been extensively characterized by other methods, crystalstructure information greatly helped to further elucidate the structure-function relationship. The Rb. sphaeroides reaction center was crystallized also by the vapor diffusion method using a solution containing 5 mg RC per mL plus heptane-triol, polyethylene glycol and LDAO. The major structural features of Rp. viridis and Rb. sphaeroides were found to be conserved, with the Rb. sphaeroides reaction center also containing L-, M- and H-subunits, with 5, 5 and 1 transmembrane helices, respectively. Except for the carotenoidless strain of Rb. sphaeroides R-26, both reaction centers contain 14 cofactors, but the cofactors are not quite the same. First, Rp. viridis contains BChl b and molecules but Rb. sphaeroides contains BChl a and Next, both and in Rb. sphaeroides are ubiquinones, while in Rp. viridis is menaquinone and a ubiquinone. There are other differences. The reaction-center crystals from Rb. sphaeroides retain both and while those from Rp. viridis lose most of their during isolation in the earlier reported studies. On the other hand, Rp. viridis reaction-center crystals still retain the two low-potential and the two high-potential c-type cytochromes, while the reaction-center of Rb. sphaeroides is known to have lost its two cytochromes during preparation. This difference in cytochrome binding strength is presumably the result of different amino-acid composition near the binding sites.

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The crystal structures of Rb. sphaeroides reaction centers from the carotenoidless strain R-26 and from the wild-type strain 2.4.1. were initially determined at 2.8 Å and 3.0 Å resolution, respectively. The structure of the Rb. sphaeroides R-26 reaction center is shown by the model in Fig. 8. Major structural features, such as a hydrophobic core containing 11 transmembrane helices from the L-, M- and Hsubunits, the approximate rotational symmetry of the L- and M-subunits, the position and orientation of cofactors, etc., are all apparently conserved. All cofactors are associated with the LM-complex. Note that the helices and their connecting segments are designated with upper- and lower-case letters, respectively, and the meaning of the underlining is the same as that for the Rp. viridis model in fig. 7. The dot in the circle slightly below the center represents the nonheme iron atom, which is approximately equidistant from the two quinone molecules.

ll.B.3. Spatial Arrangement of the Polypeptide Subunits In the following discussion, we will combine information obtained from crystallographic measurements with that from the amino-acid sequences of the three subunits from Rb. sphaeroides and Rp. viridis shown earlier in Fig. 3. Information derived from X-ray crystallography on the transmembrane helix segments for the L-, M- and H-polypeptides of Rb. sphaeroides is presented in Table 3.

In In the amino-acid sequence shown earlier in Fig. 3, the transmembrane helices as predicted by hydropathy plots are indicated by dashed lines just above and spanning the appropriate residues in the sequence, and labeled L or M followed by A to E, plus two demarcating vertical lines with residue numbers (see Table 3). It may be noted that the helical regions obtained from hydropathy plots for the three polypeptide subunits agree well with the crystallographic results. The L-subunit of the reaction center of Rb. sphaeroides contains approximately 50 fewer amino-acid residues than the M-subunit. On the other hand, the L- and M-polypeptides are similar in their each having five transmembrane helices, most of them containing 21 to 28 amino-acid residues. In fact, 216 of the M-polypeptide can almost be superimposed onto a set of corresponding of the L-polypeptide by an ~180° rotation around the principal axis of the reaction-center complex. Furthermore, most of the segments connecting the helices in both subunits are also structurally similar, some consisting of short helices themselves. The H-polypeptide [Fig. 3 (B)] is the smallest among the three polypeptides, containing only 258 amino acids. The segment of 26 amino-acid residues [I1e-12 to Arg-37] near the N-terminus forms its

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only transmembrane helix, which is approximately antiparallel to and in contact with the (double-underlined) E-helix of the M-subunit (see Figs. 7 and 8). The H-subunit helix is followed by seven consecutive charged residues and then a short disordered segment. Following this disordered region, the H-subunit forms an extended structure along the surface of the LM core near the cytoplasmic side. As the H-subunit contains no pigments or cofactors, it is not directly related to photochemical activity. Its cytoplasmic domain does appear to shield the -binding site from the cytoplasm. It has been suggested that possibly it has a role related to the electron transfer between and A structural function has also been proposed for the H-subunit in initiating the assembly of the core complex containing the reaction center and the light-harvesting complex. It is of interest to note that the green bacterium aurantiacus does not contain the H-subunit. The cofactors linked to the amino-acid residues on the L- and M-subunits might be labeled with subscripts “L” and “M.” However, this designation becomes a little complicated when it comes to the quinones, as the quinone that is buried in a pocket formed from amino-acid residues of the M-subunit is actually located in a region dominated by a branch of the L-subunit. We will therefore adopt a nomenclature which uses the subscript “A” for an L-dominated branch and “B” for an M-dominated branch, i.e., and for BChls b, and for b, and and for the two quinones. Thus menaquinone-9, although bound to the M-subunit is located at the end of an L-branch, while ubiquinone-9, is bound to the L-subunit but located near the end of an M-branch. It is to be noted that ~70% of the -binding sites in the Rp. viridis crystals appear as empty cavities on the electron-density map, as most of the was lost during isolation of the Rp. viridis reaction center. This cavity, however, can be filled with some exogenous quinone or a molecule of the inhibitors o-phenanthroline or terbutryn. The spatial relationship of the 14 cofactors to each other in the Rp. viridis reaction-center complex is presented in Fig. 9 without the background protein structure. For the sake of clarity, the phytyl and isoprenoid side chains are omitted in the figure. The primary donor, i.e., the special pair represented as and as well as the monomeric BChl-b and the molecules, are arranged on an approximate “twofold rotation symmetry axis.” This rotation axis passes between the special pair of BChl b molecules and through the nonheme iron atom. This symmetry arises from the fact that the polypeptides of the L- and M-subunits, with which these cofactors are associated, have several homologous amino-acid sequences and are therefore folded in a similar way. The tetrapyrrole rings of BChl b and on one branch can be virtually superimposed onto those of BChl b and on the other branch by a rotation of the reaction center by approximately 180°. On detailed examination, however, the overall symmetry of the cofactors is violated at several places due to local differences in the interatomic distances and interplanar angles in the two branches, differences in the phytyl chains of the BChls and the different chemical nature of the quinones and the presence of a single carotenoid (1,2-dihyroneurosporene in Rp. viridis and spheroidene in Rb. sphaeroides) near but no corresponding carotenoid near These structural differences, together with the different protein environments of the cofactors, presumably account for the difference in electron-transfer behavior of the two branches. The two BChl-b molecules located at the interface of the L- and M-subunits between the D- and Ehelices near the periplasmic side constitute the so-called “special pair,” which is the primary electron donor (abbreviated as “D”) in the bacterial reaction center. The primary donor is also referred to by its absorption wavelength, e.g., P960 in Rp. viridis and P870 in Rb. sphaeroides. We retain “P,” which has traditionally stood for the word “pigment,” since its introduction by Louis Duysens in 1952, as the symbol for the special-pair electron donor, noting its association now also with the word “pair.” Even though the two BChl molecules forming the special pair [BChl]2 belong to both branches, each individual BChl in this pair may be distinguished by the subscripts A and B and the special-pair donor P then written as

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Fig. 10 presents the values of distances between the cofactors in the reaction centers of Rb. sphaeroides and Rp. viridis obtained in a structural determination at 2.65 Å resolution as reported by Ermler, Fritzsch, Buchanan and Michel13. Some of these results will now be described. The pyrrole rings I of and are ~3.2 Å apart, but the ring centers are ~7 Å apart and thus the tetrapyrrole rings are inclined to each other by ~15°. The Mg atoms of and are each ligated to L-His 173 and M-His200, respectively, as marked by upward arrows in Fig. 3 (A). The acetyl group on ring I of forms a hydrogen bond with

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tyrosine on one side (M-Tyr208 for both species) and with histidine on the other (L-His 168 for both species). The tyrosines are in van der Waals contact with the respective rings I of both and and are likely candidates for serving as a conduit for electron transfer between the latter two (see Chapter 5 for further details). Coming off from are the “accessory” pigments, monomeric and and and followed by a quinone molecule, or The center of either monomeric BChl-b molecule is ~11 Å from that of the nearest D, and the corresponding edge-to-edge distance is ~ 4 Å. Each is ~ 10 Å from the nearest monomeric BChl and ~12 Å from the nearest quinone. The Mg atoms of and are ligated by L-His 153 and M-His 180, respectively, as marked in Fig. 3 (A). The Mg atoms of all four BChl molecules are thus penta-coordinated. The molecule, as mentioned above, is coordinated to the L-subunit, but is surrounded by two amino-acid residues from the M-subunit, M-His217 (which is also one of the ligands for Fe) and MTrp250. is linked to a site symmetrical to the site, but differs from by being nearer to the Hsubunit. This proximity of the H-subunit helix to appears to have a dramatic effect on the behavior of the two quinone molecules. Although the removal of the H-subunit does not affect the photochemical activity of the remaining LM-complex, in its absence can accept two electrons, with electron transfer to the secondary quinone being inhibited. A nonheme iron atom is located between the two quinones, but not directly attached to either. It is ligated by five amino-acid residues: two from the L-subunit (L-His 190 and L-His230), three from the M-subunit (M-His217, M-His264 and the bidendate M-Glu234), together forming a distorted octahedron about the iron. The reaction center contains one carotenoid molecule, except in the carotenoidless strain R-26 of Rb. sphaeroides. Both the spheroidene in Rb. sphaeroides and the 1,2-dihydroneurosporene in Rp. viridis assume the 15,15'-cis configuration and are located near the molecule (see Figs. 9 and 10). The protein environment around the carotenoid consists of a large number of aromatic residues, which probably impose strong steric constraints on the carotenoid, and may account for the red shift in the absorption spectrum of the carotenoid relative to that of the free carotenoid. The proximity of the carotenoid to BB suggests that the latter could serve as a conduit for the transfer of triplet-state energy from the primary donor to the carotenoid. In Rp. viridis the four hemes in the cytochrome subunit are connected to the cysteine residues in the protein via thioether bridges. The heme planes are parallel to the helical axes, but only the special bacteriochlorophyll pair and the heme of the nearest cytochrome share the same axis. The distance between this cytochrome iron atom and either Mg atom of is 21 Å. The iron-iron distance within each cytochrome pair is 14 Å while that between two cytochrome pairs is 16 Å. As expected, extracts electrons from the nearest cytochrome.

III. Summary – Reaction-Center Structure and Photochemical Reactions We now summarize in Fig. 11 the reaction-center structure and the known electron-transport reactions in purple bacteria. A simplified representation of the reaction-center and the light-harvesting complexes contained in the bacterial membrane is shown in Fig. 11 (A), followed by a column model and a cofactor model in Fig. 11 (B). The cofactor model is used to illustrate the various electron-transport steps with the associated rate constants in Fig. 11 (C), where the cofactors in the starting state (oxidized or reduced) are shown in solid black. When a cofactor first becomes reduced or oxidized, it is shown as an open symbol. We will also use this cofactor model and reaction sequence as a framework for introducing the remaining chapters throughout the section on photosynthetic bacteria.

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The next chapter will deal with the light-harvesting complexes, their structure and the mechanism of excitation-energy transfer. We will then discuss the primary electron donor (Chapter 4), a special pair of bacteriochlorophyll molecules, which is the interface between photochemistry and chemistry and the starting point for photosynthetic electron transport. We will then discuss the so-called “stable primary” electron acceptor, quinone and the secondary quinone acceptor in Chapters 5 and 6, respectively .This will be followed by a discussion in Chapter 7 of the rapid electron transport from the primary donor to and a discussion of the discovery of as the initial electron acceptor. In spite of this sequence of presentation, it should be noted that during the history of photosynthesis research, quinone was actually the earliest electron acceptor to be uncovered, chiefly as a result of lack of sufficient time resolution in the instruments used in the earlier work. With only micro- or milli-second time resolution, reactions involving the earlier electron acceptors could of course simply not be detected. The section on photosynthetic bacteria will be concluded with a discussion of the electron donor to the oxidized special pair, namely, the cytochrome, in Chapter 10. The discussion of green photosynthetic bacteria will be presented in Chapters 8 and 9, respectively, dealing with the light-harvesting complex, chlorosomes, and the reaction centers of green bacteria.

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For further reading R1. RE Blankenship, MT Madigan and CE Bauer (eds) Anoxygenic Photosynthetic Bacteria, 1995. Kluwer R2. J Deisenhofer and H Michel (1989) The photosynthetic reaction centre from the purple bacterium Rhodopseudomonas viridis (Nobel Lecture). The EMBO J 8:2149-2169 R3. DC Rees, H Komiya, TO Yeates, JP Allen and G Feher (1989) The Bacterial photosynthetic reaction center as a model for membrane proteins. Annu Rev Biochemistry 58: 607-633 R4. JR Norris and A Schiffer (1990) Photosynthetic reaction centers in bacteria. C&ENews 68 (31): 22-37 R5. CRD Lancaster and H Michel (1995) The structures of photosynthetic reaction centers from purple bacteria as revealed by X-ray crystallography. In: Anoxygenic Photosynthetic Bacteria (see R1) pp 503-526 R6. G Drews and JR Golecki (1995) Structure, molecular organization, and biosynthesis of membranes of purple bacteria. In: Anoxygenic Photosynthetic Bacteria (see R1) pp 231-257 R7. A Vermeglio, P Joliot and A Joliot (1995) Organization of electron transfer components and supercomplexes. In: Anoxygenic Photosynthetic Bacteria (see R1) pp 279-295 R8. G Feher (1998) Three decades of research in bacterial photosynthesis and the road leading to it: A personal account. Photosynthesis Research 55: 1-40

References 1. 2. 3. 4. 5.

6. 7. 8.

9. 10.

11.

12. 13. 14.

DW Reed and RK Clayton (1968) Isolation of a reaction center fraction from Rhodopseudomonas sphaeroides. Biochem Biophys Res Commun 30: 471-475 RK Clayton and RT Wang (1971) Photochemical reaction centers from Rhodopseudomonas sphaeroides. In: A San Pietro (ed) Methods in Enzymology, Vol 23A, pp 696-704. Acad Press G Feher and MY Okamura (1978) Chemical composition and properties of reaction centers. In: RK Clayton and WR Sistrom (eds) The Photosynthetic Bacteria, pp 349-386. Plenum G Gingras (1978) A comprehensive review of photochemical reaction center preparations from photosynthetic bacteria. In: RK Clayton and WR Sistrom (eds) The Photosynthetic Bacteria, pp 105-132. Plenum C Hager-Braun, A Riedel, DL Xie, R Zimmermann, N Nelson and G Hauska (1995) Characterization of the photosynthetic reaction centre from Chlorobia. In: P Mathis (ed) Photosynthesis: From Light to Biosphere, Vol II, pp 171-174. Kluwer J Kyte and RF Doolittle (1982) A simple method for displaying the hydropathic character of a protein. J Mol Biol 157: 105-132 H Michel (1982) Three-dimensional crystals of a membrane protein complex. The photosynthetic reaction centre from Rhodopseudomonas viridis. J Mol Biol 158: 567-572 J Deisenhofer, O Epp, K Miki, R Huber and H Michel (1984) X-ray structure analysis of a membrane protein complex. Electron density maps at 3 ¯ resolution and a model of the chromophores of the photosynthetic reaction center from Rhodopseudomonas viridis. J Mol Biol 180: 385-398 JP Allen and G Feher (1984) Crystallization of reaction center from Rhodopseudomonas sphaeroides: preliminary characterization. Proc Nat Acad Sci, USA 81: 4795-4799 C-H Chang, M Schiffer, D Tiede, U Smith and J Norris (1985) Characterization of bacterial photosynthetic reaction center crystals from Rhodopseudomonas sphaeroides R-26 by X-ray diffraction. J Mol Biol 186: 201 203 B Arnoux, JF Gaucher, A Ducruix and F Reiss-Husson (1995) Structure of the photochemical centre of a spheroidene-containing purple bacterium, Rhodobacter spheroides Y, at 3 ¯ resolution. Acta Crystallog, Section D, 51: 368-379 J Deisenhofer, O Epp, I Sinning and H Michel (1995) Crystallographic refinement at 2.3 ¯ resolution and refined model of the photosynthetic reaction center from Rhodopseudomonas viridis. J Mol Biol 246: 429-457 J Breton and E Nabedryk (1987) Pigment and protein organization in reaction center and antenna complexes. In: J Barber (ed) The Light Reactions, pp 159-195. Elsevier U Ermler, G Fritzsch, SK Buchanan and H Michel (1994) Structure of the photosynthetic reaction centre from Rhodobacter sphaeroides at 2.65 ¯ resolution: cofactors and protein-cofactor interactions. Structure 2: 925936

Chapter 3 Light-Harvesting Pigment-Protein Complexes of Photosynthetic Bacteria I.

The Light-Harvesting Pigment-Protein Complexes A. Isolation and Spectral Properties B. Protein structure II. An Early Model for the Light-Harvesting Pigment-Protein Complex III. Crystal Structure of the Light-Harvesting Complexes of Purple Photosynthetic Bacteria A. LH2 (B800-850) of Rhodopseudomonas acidophila B. LH2 (B800-850) of Rhodospirillum molischianum C. LH1 (B890) of Rhodospirillum rubrum D. LH1 (B875) of Rhodobacter sphaeroides a Computational Model E. The Dimeric LH1•RC Supercomplex of Rhodobacter sphaeroides IV. Structure and Function of the Bacterial Photosynthetic Apparatus

66 66 68 69 71 71 75 77 78 78 81

The photosynthetic apparatus of purple bacteria consists in part of a reaction center surrounded by several light-harvesting complexes, as shown schematically in Fig. 1. The light-harvesting complexes B800-820 and B800-850 (LH2) and B875 (LH1) absorb light energy over a moderately wide spectral region in the near infrared (as well as in the blue region); the numbers in the names of the complexes refer to the wavelength, in nm, of characteristic absorption peaks. This energy is transferred down the B800/B820/B850/B875/B890 excitation-energy gradient, finally reaching the reaction center where it initiates photochemical charge separation and electron transport. This chapter discusses the isolation, structure and properties of these light-harvesting complexes. The role of carotenoids in the bacterial light-harvesting complex will be discussed briefly but in more detail later in Chapter 13. General aspects of energy transfer involving bacteriochlorophylls and carotenoids was first introduced in Chapter 1.

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I. The Light-harvesting Pigment-Protein Complexes

I.A. Isolation and Spectral Properties The near-infrared absorption spectra of purple photosynthetic bacterial cells as well as their chromatophore membranes show absorption bands near 800, 820, 850 and 870-890 nm, as illustrated in Fig. 2 (B) by the spectrum of Chromatium-vinosum chromatophores, where most of the absorption bands originate from the BChl molecules present in the membrane. The BChl-a molecules are present in three lightharvesting BChl-protein complexes: B875 with an absorption peak between 870 and 880 nm, B800-850 with peaks at 800 and 850 nm, and B800-820 with peaks at 800 and 820 nm. Here the letter B refers to “bulk pigment,” and the numbers refers to the wavelengths of absorption peak of the complex in the near infrared. There are two types of light-harvesting complexes: the longer wavelength-absorbing “core” (or “inner,” or “proximal”) antenna, also known as “light-harvesting complex 1” or “LH1,” e. g., B875 in Rhodobacter sphaeroides or B890 in Rhodospirillum rubrum, and the shorter wavelength absorbing “peripheral” or “distal” complexes B800-850 and B800-820 (collectively known as “light-harvesting complex 2” or “LH2”). The core antenna complex is present in all photosynthetic bacteria and is intimately associated with the reaction center. As illustrated in Table 1, different photosynthetic bacteria can vary in their makeup of antenna complexes. For example, Rhodospirillum rubrum and the BChl b-containing Rp. viridis contain only the core antennae, B890 and B1015, respectively. It might be noted that B1015 is the longest wavelength absorbing antenna pigment yet known. Rhodobacter sphaeroides and Rhodobacter capsulata, on the other hand, contain both the core antenna B875 and the peripheral antenna B800-850. Chromatium vinosum and Rhodpseudomonas acidophila contain the core antenna and two additional peripheral antenna complexes, B800-850 and B800-820.

The antenna complexes transfer absorbed light energy along a gradient of excitation energy extending from the outermost, shortest wavelength complex in toward the reaction center. Unlike the core antenna, which is present in a fixed stoichiometry relative to the reaction center, the relative number of chlorophyll molecules in the peripheral antennae may vary as a result of adaptation to the environmental conditions such as light intensity or temperature1. Under growth condition of high light intensity, for example, the peripheral antennae are present in smaller amounts, but in larger amounts when grown under low light intensity. This adaptability allows the bacteria to optimize their light-harvesting capacity with respect to the organism’s ability to use this energy in the subsequent dark reactions, according to the amount of light available by regulating the size of their photosynthetic unit according to the amount of light available.

Chapter 3

Light-Harvesting Pigment-Protein Complexes of Photosynthetic Bacteria

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When BChl a is dissolved in an organic solvent it shows a single absorption band at 772 nm in the farred region, but when BChl a is bound to a protein its absorption band is red-shifted to 800-900 nm as a result of the influence of its protein (and lipid) environment. The absorption peak of BChl b in an organic solvent absorbs at 790 nm, but when bound to a protein in an antenna complex its absorption band also shifts, in this case to 1015 nm. When the pigment-protein complex is denatured in some way, the redshifted absorption band is seen again at the original shorter wavelength. Since the bacteriochlorophyll present in the light-harvesting complex accounts for the majority of all the bacterial pigments, its absorption bands can readily be identified even in the spectrum of the unfractionated membrane. On the other hand, the pigments belonging to the reaction center amount to only ~1% of the total BChl and its absorption is often masked by the bulk pigments. The BChl a present in the reaction center may be identified however in a purified reaction-center preparation isolated from the chromatophore membrane. This may be illustrated with Chromatium vinosum following fractionation and isolation of the reaction-center complex and the three antenna complexes from the chromatophore membrane. Fig. 2 (B) shows the absorption spectrum of the unfractionated Chromatium chro-

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matophores, while Fig. 2 (A) shows that ofthe isolated Chromatium reaction-center complex. The solidline spectrum in Fig. 2 (A) represents the sample in the reduced state produced by addition of dithionite and the dashed-line spectrum the sample in the oxidized state by addition of ferricyanide or by illumination. The three spectra in Fig. 2 (C) are those ofthe three different light-harvesting, bacteriochlorophyllcarotenoid-protein complexes. The four component spectra in Figs. 2 (A) and (C) should sum up to that of the unfractionated membrane in Fig. 2 (B). Spectrum analysis of the chromatophores shows that the 883-nm band arises from the B890 complex, the 840-nm band from the B800-850 complex and the 804nm band from a combination of the B800-850 and B800-820 complexes. The absorption band in the 480-nm region in the chromatophore membrane and in the three light-harvesting complexes is mainly due to carotenoid. Also, all BChls have a so-called “Soret” band near 371 nm. Note that in the spectrum of the B800-850 complex from C. vinosum, the 803 nm absorption band is only just slightly more intense than the band at 846 nm. On the other hand, the B800-850 complex isolated from Rhodobacter sphaeroides or Rhodopseudomonas capsulata has an 850-nm band about 1.5 times higher than the 800-nm band (not shown here). When the B800-850 complex of a given species is found to have a high 800/850 ratio, it has been shown that there is always a B800-820 BChl-protein complex present [see Fig. 2 (C), left]. B800-820 is spectrally rather similar to B800-850, suggesting the former complex may be a variant of the latter.

I.B. Protein Structure The “minimum” structural unit of each of the three light-harvesting pigment-protein complexes consists of two BChl molecules and one carotenoid molecule, each noncovalently bound to two low molecular-weight (5-7 kDa) polypeptides designated as and In the core antenna complex B1015 of Rp. viridis and other BChl b-containing species, however, there is also a The amino-acid sequences of the and of the B870 and B800-850 complexes from a number of photosynthetic bacteria is shown in Fig. 3. Each of the and contains about

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50-60 amino acids. Sequence analysis shows that all the and have three domains. The polar N-terminal domain is located at the cytoplasm. The center portion ofthe polypeptide chain is made up of ~20 hydrophobic amino acids which form an spanning the membrane. The polar C-terminal domain is located near the periplasmic side of the membrane. The has a single conserved histidine in the hydrophobic domain near the C-terminus (marked “[1]” in Fig. 3), while the has a pair of conserved histidines, one in the hydrophobic domain near the C-terminus and the other in the N-terminal domain (marked “[2]” in Fig. 3). Each ofthese conserved histidines is thought to serve as a ligand to a Mg atom of a BChl molecule, as suggested by resonance Raman studies. Various other conserved amino acids or their clusters may have specific functional roles with regard to pigment binding, protein conformation or, indirectly, spectral properties. For instance, the conserved alanine (marked “[3]”), which is situated one helical turn away from the conserved histidine residue, “[1],” has been postulated as an additional interaction site for the BChl a heterodimer of the reaction center. It may be noted that the N-terminal domain of the is shorter than that of the while the C-terminal domain of the is correspondingly longer. These differences may be of significance to the membrane surface organization. Note also that the and of each complex show only 7-12% sequence homology, indicating a separation of the and by gene duplication during the course of evolution. On the other hand, the (and polypeptides from different light-harvesting complexes of the same organism show a relatively high degree of sequence homology. The (and polypeptides from the same antenna complex but from different organisms also show a high sequence homology, suggesting a species relationship between antenna systems.

II. An Early Model for the Light-Harvesting Pigment-Protein Complex On the basis of the primary-structure information discussed above, and the known, 1:1 stoichiometry of the and developed a model for the light-harvesting BChl-protein complexes, as shown in Fig. 4 (A). In this model the two polypeptides span the chromatophore membrane with their hydrophobic segments consisting of ~20 amino acids located in the hydrocarbon-tail region of the lipid-bilayer membrane. The polar N- and C-termini are located on the cytoplasmic and periplasmic sides, respectively. The hydrophobic amino acids in this model are present as an in accord with 2 the finding of a high helical content by circular dichroism . In the model for B890, B850 or B1015, each of the binds a BChl-a molecule with the conserved histidine residue [shown as bold H in Fig. 4 (A)] near the C-terminus. In B800-850, each histidine near the C-terminus binds a BChl-a molecule that absorbs at 850 nm while a second conserved histidine near the N-terminus of the presents one more binding site for yet another BChl a, the one absorbing at 800 nm. In the B890 and B800-850 complexes, strong CD bands appear in the 890- and 850-nm regions, respectively, indicating the presence of a pair of interacting BChl molecules3. The 800-nm absorptionband region shows no optical activity and is therefore assigned to a single BChl-a molecule. The analogue of the B890 complex in Rhodobacter sphaeroides, B875, contains two BChl a and two carotenoid molecules, but the CD spectrum near 880 nm is not very intense, indicating only a weak interaction between the BChl-a molecules. The counterpart of B875 in the carotenoidless mutant of Rhodobacter sphaeroides, namely, B860, also contains two BChl-a molecules associated with the two polypeptides. In this model the is formed predominantly by an interaction of the amino-acid side chains near the cytoplasmic N-terminus. Fig. 4 (B) shows the in a tight packing arrangement, with the axes of the two helices at angle of ~30° to each other (note that only the hydrophobic domains are shown). The figure also shows that planes of the two BChl-a molecules forming the primary

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donor pair are approximately perpendicular to the membrane plane. The is asymmetric, with the BChl molecules located near the periplasmic side, and the cytoplasmic side providing a binding site for the polypeptides. The may thus be considered as the minimum unit of the lightharvesting complex, in terms of spectral properties. The particular location of the BChl pair forms

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the basis for two-dimensional energy transfer from the antenna complexes to the special BChl pair ofthe reaction center, which is also located near the periplasmic region. The through their amino-acid side groups, can further interact to form larger aggregates of the antenna complex. One hypothetical model for B800-850 is a hexamer (i.e., in which six twisted aggregate into a larger assembly, with the hatched areas in Fig. 4 (C) representing 4 ofthe 12 BChl molecules. It should be mentioned that Scherz and Parson4, using spectroscopic data, have also developed a model for the BChl-protein antenna complex. In their model, 2 heterodimers are assumed to be coupled to form as the minimum unit of the antenna complex. In the late 1980s, several bacterial antenna complexes were crystallized5-11, but none appeared suitable for high-resolution, X-ray crystallography. Among crystals prepared from an antenna complex, only that of B800-850 ofRps. acidophila has been studied. Results from preliminary crystallographic studies were interpreted as suggesting that an antenna complex contained three bundles of units arranged as shown in Fig. 4 (D). Such an antenna complex would contain 6 and 6 18 BChl-a and 9 carotenoids. The distance between each helical pair is 10.5 Å [marked “a” in Fig. 4 (D)], the diagonal packing distance is 14.2 Å (marked “b”), and the distance between two bundles is 24.0 Å (marked “c”). This model would also yield a molecular weight of 84 kDa for the intact complex, in agreement with reported values obtained by gel electrophoresis and by ultracentrifugation. However, as seen below in section IV, recent X-ray crystallographic measurements with suitable crystals of this light-harvesting complex have revealed an exquisite three-dimensional structure apparently quite different from those shown in Fig. 4.

III. Crystal Structure of the Light-Harvesting Complexes of Purple Photosynthetic Bacteria Many difficulties needed to be overcome in the pursuit of a crystal of satisfactory quality, but once a suitable crystal became available, the successful determination of the three-dimensional structure ofthe bacterial light-harvesting complex quickly became a reality. The first such report was for the B800-850 complex (LH2) of Rp. acidophila which had previously been extensively studied by standard methods and its crystallization intensively pursued7, 8.

III.A. LH2 (B800-850) of Rhodopseudomonas Acidophila In 1995, the three-dimensional structure of the light-harvesting complex B800-850 (LH2) of the photosynthetic purple bacterium, Rp. acidophila strain 10050, was determined by McDermott, Prince, Freer, Hawthornthwaite-Lawless, Papiz, Cogdell and Issacs12 by X-ray crystallography at a resolution of 2.5 Å. Contrary to the earlier model discussed in the previous section, analysis ofthe X-ray data showed that the Rp. acidophila B800-850 complex consists of 9 and 9 27 BChl-a molecules, 13-14 carotenoids, and 22 water molecules. Two cross-sectional views of the light-harvesting complex in the direction ofthe membrane plane but at two different depths along the length ofthe polypeptide, one near the periplasmic side and the other near the cytoplasmic side, are shown in Fig. 5 (A) and (B), respectively. Each subunit is associated with 3 BChl-a molecules and one carotenoid. The and consisting of 53 and 41 amino acids, respectively, are presumed to span the membrane bilayer. The polypeptides and BChl-a molecules in the cross-sectional views are labeled on the right side of Fig. 5 (A) and (B) [carotenoids are not shown]. To facilitate discussion, the helical portions of the and chains are shown with their residues labeled in Fig. 5 (C). The N-termini are located on the cytoplasmic side and the C-termini on the periplasmic side.

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As shown in Fig. 5 (A), the 9 helical are arranged in the form of an inner cylinder with a 36-Å diameter, while the 9 form a concentric outer cylinder with a 68-Å diameter. Each is parallel to the 9-fold axis to within an experimental error of ±2°, whereas each is inclined by 15° relative to this axis. The transmembrane helices formed by residues 11 to 36 [see Fig. 5 (C)] of both the and (PP) are anchored to amphipathic helical tails at the N- and C-termini. The following hydrophobic residues are marked with solid dots in Fig. 5 (C): Ile-6, Trp-7, Val-9, Trp-40, Phe-41, Tyr-44, Trp-45; and Ala-1, Leu-3, Ala-5, Pro-38, Trp-39, Leu-40. have an anchoring effect in the membrane, while the hydrophilic residues lie on the cytoplasmic and periplasmic membrane surfaces, consonant with the bilayer lipid membrane. The 27 BChl-a molecules are distributed into two groups. The 18 BChl-a molecules of the first group, located near the periplasmic end of the complex, are ligated through their central Mg atoms to the conserved residues His-31 and His-30 of the and respectively. These and

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are pointing outward and inward, respectively, i.e., toward the interior of the annular space between the and Thus a complete ring of 18 overlapping BChl-a molecules is formed within the annular space. The planes of the BChl-a macrocycles are perpendicular to the membrane, with the center of each macrocycle ~10 Å from the periplasmic membrane surface. The phytol chains point toward the hydrophobic core of the assembly. These overlapping BChl-a molecules are strongly exciton-coupled, and are responsible for the 850-nm absorption of the B800-850 light-harvesting complex. The second group of 9 BChl-a molecules is near the cytoplasmic end, with each sandwiched in between the helices [see Fig. 5 (B)] and with their macrocycle planes parallel to the membrane. Each of these pigment molecules is kept in position by ligation of its central Mg atom to the formyl carbonyl oxygen of the formylmethionine-1 residue at the N-terminal position of the Note that this amino acid in most of the published literature is reported as a methionine, but recent X-ray work found it to be formylmethionine (“fMet” or “fM”), buried ~9 Å from the cytoplasmic membrane surface. The ligated BChl-a molecules are ~11 Å from the cytoplasmic surface while the Mg-toMg distance between neighboring BChl-a molecules is 21 Å. It is these 9, largely non-interacting (i.e., monomeric) BChl-a molecules that make up the 800 nm-absorbing component of the B800-850 complex. The phytol chains of these 9 BChl-a molecules on the cytoplasmic side are directed into the hydrophobic core of the assembly and are in van der Waals contact with each other, and the polypeptide residues also intertwined, with the phytol chains of the 18 B850 BChl-a molecules on the periplasmic side, which themselves are coordinated to the and The Mg-to-Mg distance between the two rings of BChl-a molecules in B800 and B850 is 17.6 Å. Two detailed views of the B800-850 complex, one normal to and one along the membrane plane, are shown in Fig. 6 (A) and (B), respectively. Here the and BChl-a and carotenoids [in (B) only] are shown. The placement and orientation of the polypeptide and pigment molecules can be readily discerned. Corresponding color figures of the Rp. acidophila LH2 complex are shown as Color plates 3 (A) and (B) in the color-plate section near the beginning of the book. It should be mentioned that more than 10 years before the X-ray diffraction studies the orientation of the BChl-a molecules of B800 and B850 was correctly deduced by Kramer, van Grondelle, Hunter, Westerhuis and Amesz13 from fluorescence-polarization studies. These authors also determined the ratio of the BChl-a molecules in B850 to B800 to be 2, but their proposed a model consisting of only 4 B850-BChl a, 2 B800-BChl a and 3 carotenoid molecules in the complex. Also noteworthy is the prediction made sometime ago by Monger and Parson14, Sauer and Austin3, Miller15, l6 and Stark, Kühlbrandt, Wildhaber, Wehrli and Mühlethaler17 that the bacterial reaction center is surrounded by the core antenna complex LH1 (B890). However, the assuming of an analogous cylindrical shape by LH2 (B800-850), given its homology with LH1, was not anticipated. As shown in Fig. 4 (D) above, LH2 had been assumed in one model to be a cyclic trimer of units held together by hydrophobic interactions between the polypeptide helices, in some ways similar to the current model. The environments of the macrocycles of two groups of BChl-a molecules in B800-850 are distinctly different. That for the 18 BChl-a molecules of B850 is predominantly hydrophobic, as they are in contact with various non-polar amino-acid residues from both the and (marked with small dots and numbered in Fig. 5 (C)]. On the other hand, the 9 B800 BChl-a molecules are in a relatively polar environment; each is in contact with His-12 from the the formyl group on fMet on the and a water molecule near His-12. There is also a strong hydrogen bond between Arg-20 of the and the 2-acetyl carbonyl oxygen of the BChl a. The X-ray structural results can therefore account for the different optical properties of the BChl-a molecules in B800 and B850 in terms of their chemical environment.

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The X-ray crystal-structure determination revealed a lack of any direct, helix-helix interaction between the and within the hydrophobic core, as had been expected on the basis of earlier models [see Fig. 4 (B) and (C) above]. The association of the fMet-1 residue at the N-terminal of each to a B800 BChl-a molecule prevents direct interaction between radially paired and Interactions between the membrane-core helices are mediated only via pigment molecules or buried water molecules. The only polypeptide interaction occurs at the N- and C-terminal tails. In the C-terminal tail the large aromatic residues and contribute to binding between polypeptides via hydrogen bonds and hydrophobic interactions [see Fig. 5 (C)]. Each molecule of the carotenoid rhodopin glucoside [see Fig. 5 (D)] in the B800-850 light-harvesting complex is in its extended conformation and able to span the width of the membrane. The glucosyl ring is associated with the via Lys-5 and Thr-8 [marked with an “x” and numbered in Fig. 5 (C)]. Along the carotenoid chain, van der Waals contact (CP29, with the two core antenna complexes CP43 and CP47 containing Chl a but containing no Chl b at all. For reference purpose, we list in the table at left the names used in this book and the alternative nomenclature used in photosynthesis literature1.

I. A. The Core Antenna Complexes CP47 and CP43 The core antennas CP47 and CP43 are Chl-a proteins that participate in both light harvesting and transfer of energy to the reaction center of photosystem II, with CP47 interacting more directly and more strongly with the reaction center than CP43. While the number of the other antenna complexes per PS-II reaction center is somewhat variable, there is always one CP43 and one CP47 complex per reaction center, i.e., per D1/D2 heterodimer. CP47 and CP43 migrate on SDS-PAGE with apparent molecular weights of 45-51 kDa and 40-45 kDa, representing, in reasonable agreement with their molecular masses (51 and 44 kDa) derived from the nucleotide sequences of their respective genes pscB and pscC. Each polypeptide contains 10-12 conserved histidine residues that bind the chlorophyll molecules via ligation to the Mg atoms. In addition to light harvesting, these complexes may also serve to anchor proteins in certain PS-II reaction-center complexes. CP47 and CP43 consist of ~190 and ~130 amino acids, respectively. Hydropathy-plot analysis of their amino-acid sequences shows that both polypeptides have six transmembrane each polypeptide having a large hydrophilic domain between the last two helices. These large loops are exposed to the inner thylakoid surface on the lumen side and it has been suggested that they may serve as anchors for the three extrinsic proteins and the manganese cluster that are involved in oxygen evolution [see Fig. 1 (B)].

I.B. The Minor Chl a/b Binding Complexes CP29, CP26 and CP24 The three minor types of Chl-a/b binding proteins are somewhat more distant from the reaction center, with CP29 and CP26 slightly closer to the reaction center than CP24, as determined by freeze-etch EM experiments and as indicated in Fig. 1 (B). Each of the three minor Chl a/b complexes binds

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~5% of the total PS-II chlorophylls. These light-harvesting complexes, also collectively called the “accessory chlorophyll proteins” or “ACP,” are encoded by nuclear genes [see table on previous page]. The intermediate location of these complexes suggest they may have a role as linkers, or anchors, for the peripheral LHC II. Unlike LHC II, however, these minor Chl a/b complexes do not become phosphorylated under high light conditions (see Section III below), which is consistent with the notion that they are more closely associated with the inner core antenna complex than is LHC II.

I. C. The major Chl a/b Binding Protein Complex LHC II On a chlorophyll basis, the Chl a/b binding protein LHC II is the major component of the thylakoid membrane and, though located farthest from the reaction center, is the major light-harvesting antenna of photosystem II. In addition to harvesting light, LHC II is also known to play a role in membrane stacking, which has consequences in the regulation of the distribution of energy between the two photosystems (see Section III below).

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LHC II was discovered more than thirty years ago by Ogawa, Obata and Shibata2, and independently by Thornber, Smith and Bailey3, as a green band on the SDS-gel of detergent-treated spinach chloroplasts, and originally designated as “CPII.” The literature on LHC II has now become voluminous; there are probably more papers published on LHC II than on any other thylakoid component. Each monomer subunit of a LHC II [see Fig. 1 (A)] has a molecular mass of ~25 kDa and contains 15 Chl molecules – 8 Chl a and 7 Chl b – plus two xanthophyll molecules called the luteins. The LHC-II complexes contain about half of the total PS-I and PS-II chlorophyll and ~30% of the thylakoid protein, making it probably the most abundant membrane protein of green plants. LHC II is heterogeneous, consisting of two kinds of polypeptides with molecular masses of 25-kDa and 27-kDa, as revealed by SDS-PAGE. This is consistent with the finding that LHC II proteins are encoded by multiple nuclear genes. As discussed below in Section II, Kühlbrandt and coworkersR3 have recently determined the detailed structure of the LHC II antenna complex by electron crystallography. The LHC II antenna functionally exists as either an inner or a more peripheral component. An inner (“i,” also called “bound”) LHC II contains exclusively the 27-kDa apoprotein and is located closer to the PS-II reaction center. The other, more peripheral or outer complex (“o,” also called “mobile”) contains

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predominantly the 25-kDa apoprotein. The outer, 25-kDa antenna complex can become detached from PS II upon phosphorylation (see Section III below). The LHC-II polypeptide is an integral membrane protein. A hydropathy plot of the amino-acid sequence of pea LHC II, as shown in Fig. 2 (A), suggests the presence of three hydrophobic transmembrane a-helices. The regions of the presumed, helix-forming residues are marked by horizontal bars in the figure. A schematic sketch of the conformation of the amino-acid backbone of LHC II from Lemna gibba (an aquatic monocot) is shown in Fig. 2 (B). This conformation is supported by results from mutagenesis studies and chemical-labeling experiments and also subsequently by high-resolution electron microscopy and by the even more recent structure determination by electron crystallography.

II. Crystal Structure of LHC II In 1987, Kühlbrandt4 succeeded in preparing large, two-dimensional crystals of LHC II. Subsequently Kühlbrandt and Wang5 applied the rather new technique of electron crystallography, developed earlier by Henderson and Unwin6 for determining the structure of bacteriorhodopsin, to examine the structure of LHC II at a resolution of 6 Å. Their work revealed the presence of three transmembrane in LHC II (see topography in Fig. 3), consistent with the predictions based on the hydropathy plots shown in Fig. 2. Helices A and B were found to be ~46 Å and ~49 Å long, respectively, corresponding to 31 and 33 amino-acid residues, i.e., substantially greater than the 20-22 predicted in Fig. 2. The helices A and B come within ~10 Å of each other and their axes are inclined at 25° and 31°, respectively, relative to the membrane normal, but in opposite directions, i.e., rotated about a mutually perpendicular axis, forming a “supercoil,” as shown in Fig. 3. Helix C is shorter, ~30 Å long, corresponding to ~20 amino acids, and its axis is inclined ~30° relative to the membrane normal. Counting from the N-terminus, the amino-acid chain follow the sequence B-C-A.

Although the information obtained at 6 Å resolution was insufficient to locate the porphyrin rings of chlorophyll with precision, it appears that it could be concluded that the axes of the porphyrin planes are probably oriented at least approximately perpendicular to the membrane plane and the center-to-center distances between nearest neighbor chlorophylls are between 9 and 14 Å. Also, the binding state of the 15 chlorophyll molecules could not be established since at the time only three histidines, the usual ligands for the Mg-atom of chlorophylls, were known to be contained by the polypeptide.

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II. A. Polypeptides of the LHC II Subunit A refined structure determination of LHC II at 3.4 Å resolution by Kühlbrandt, Wang and Fujiyoshi7 in 1994 revealed additional features in the light-harvesting complex. The higher resolution was afforded by the use of an electron microscope with a specimen stage cooled with liquid helium to 4.2 K. A side view of the LHC II monomer structure and the corresponding representation of the amino-acid sequence are shown in Figs. 4 (A) and (B), respectively. The B-C-A polypeptide sequence is the same as that for the polypeptide in Fig. 3. It was now found that helix-B is 51 Å long and includes 35 residues (from Pro65 to Gly-89) in 9.5 turns and helix-A is 43 Å long and includes 30 residues (from Pro-170 to Ile-199) in 8 turns [see Fig. 4 (B)]. Both A- and B-helices start with proline and are inclined 32° relative to the membrane normal in opposite directions. Helix-C is 31 Å long and includes 21 residues (from Ser-123 to Ile-143) in 5.5 turns, but inclined only 9°, not 30°. The A- and B-helices were now determined to be 8.5 Å apart, and that the two helices intertwine to form a left-handed supercoil. An additional helix (helix D), containing only 10 residues (from Pro-205 to Ala-214) in ~3 turns, was found to be located at the interface between the hydrophobic interior and the thylakoid lumen. The secondary structure of two areas of the membrane-spanning protein has not yet been well resolved, one being the N-terminus which contains the regulatory phosphorylation site (see section III below) and the other undefined region being the loop between helices B and C [see Figs. 4 (A) and 5 (A)]. About 36% of the total number of 232 amino-acid residues make up the transmembrane helices; the remainder are at or near the stromal and lumenal surfaces.

II.B. The Chlorophyll Molecules Although biochemical analysis showed the presence of 14 chlorophyll molecules per monomer of the crystalline LHC II preparation, 8 Chl a and 6 Chl b, their electron-density map could only account for 12. The three porphyrin planes closest to each of the two luteins are within 3.7-4.1 Å. Since the 3.4 Å resolution is still not enough to differentiate between Chl a and Chl b, Kühlbrandt and coworkers made a reasonable guess and assigned these six porphyrins to Chl a molecules and the other porphyrins to Chl b molecules. Their reasoning was based on the known fact that Chl b transfers electronic excitation energy to Chl a very rapidly (and not vice versa) and that carotenoids are generally found to quench Chl a triplets rather than Chl b triplets. As seen in Figs. 4 (A) and (B) and the table to its right, and in Fig. 5 (B), Chl a1, Chl a2 and Chl a3 (abbreviated as a1, a2 and a3) are in van der Waals contact with lutein 1 (abbreviated as L1), and Chl a4, Chl a5 and Chl a6 [also Chl a7] (a4, a5, a6 and a7) are in van der Waals contact with lutein 2 (L2). The remaining porphyrins are assigned to Chls b and Chl a8. The binding mode of the chlorophyll molecules in the light-harvesting complex has been identified for only 8 of the 12 chlorophylls. These chlorophylls are attached to the polypeptide through coordination of their central Mg-atoms either to the polar amino-acid side chains or the main-chain carbonyls in the hydrophobic interior of the complex. Only 2 of the 3 known histidines appear to serve as ligands: His-68 to a5 and His-212 (in helix D) to b3 [see Fig. 4 (B)]. Three amide side groups also serve as ligands: Gln-131 to b6, Asn-183 to a2 and Gln-197 to a3. Three charge-compensated glutamates, which form ion pairs with arginines either from the same helix [Glu-139/Arg-142] or from different helices [Glu-65/Arg-185 and Glu-180/Arg-70], bind b5, a4 and a1, respectively [see Fig. 4 (B)]. Interestingly, through inter-helical ion-pair formation, a pair of glutamates can apparently serve the additional function of binding helices A and B together. It seems likely that Chl a6 (or possibly also b1, b2 and b7) is ligated to the peptide carbonyl of Gly-78, with the possible intervention of a water molecule.

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II. C. The Carotenoid Lutein As previously noted, each LHC-II monomer contains two lutein molecules [see Figs. 4 (A) and Fig. 5 (B)]. These two all-trans polyene chains, each 30 Å long, are in the highly hydrophobic environment of helices A and B, and are inclined at an angle of 50° relative to the membrane normal. The shortest dis-

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tance between the two luteins is ~11 Å, the two luteins forming an X-like brace in the center of the polypeptide complex. As many of the amino-acid residues are used for ligating the pigment molecules present in the complex, the cross brace formed by the two luteins can help augment the structural rigidity of the polypeptide complex by linking the loops exposed on the opposite membrane surfaces. It is known that reconstitution of LHC II requires not only Chls a and b but, unexpectedly, xanthophylls as well. This can be explained by the finding that the two lutein molecules not only have a light-harvesting role but also an essential structural role in the assembly of the LHC II complex. The location of the carotenoid is also consistent with their role as a quencher of chlorophyll triplet which would otherwise lead to the formation of photoreactive singlet oxygen that can damage the thylakoid membrane. The structural features of the pigments can be further visualized from the stereo view of the LHC II complex in Fig. 5 (A) and that of the pigment molecules alone in Fig. 5 (B).

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II.D. The LHC II Trimer LHC II readily forms trimers, as found in vitro and in situ. The trimer configuration of LHC II, either in detergent-solubilized solution, as two-dimensional crystals, or in the native, functional thylakoid membrane has been determined by various physical methods, including analytical ultracentrifugation, circular-dichroism spectroscopy, and electron microscopy in conjunction with image analysis. The suggestion made earlier in Chapter 3 that trimer configuration of the bacterial light-harvesting complex may be the most suitable configuration for the purpose of light harvesting, probably also applies to LHC II in green-plant photosystem II. A side view of the LHC II monomer and a side and top view of the LHC II trimer are shown in Fig. 6. The two views of the trimer in Fig. 6 bottom shows the trimer to be a compact cylindrical unit, ~70 Å in diameter and 50 Å high. The stromal surface appears very flat, whereas the lumenal surface appears somewhat corrugated. The chlorophyll molecules appear arranged in two layers near the stromal and

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lumenal surfaces, each layer connected by the elongated carotenoid molecules. The top view (from the stromal side) in Fig. 6, bottom right, shows the left-handed supercoil formed by the central pair of helices and surrounded by chlorophyll molecules, with the Chl b molecules situated further away than the Chl a molecules [cf. Fig. 4 (B)], at the two ends of the supercoil. The two carotenoid molecules are located in the grooves of the supercoil. Contact between the monomers is mediated by the three helices in the center of the trimer and by the carboxyl- and amino-terminal loops. The shortest chlorophyll-tochlorophyll distances across adjacent subunit boundaries runs from 16 to 19 Å. These distances are longer than those within the subunit monomer, but still close enough for efficient and rapid energy transfer. Helix C, whose axis is almost parallel to the membrane normal (see top view in Fig. 6), and the N-terminal loop appear to act as spacers to prevent any closer approach between chlorophylls of different subunits.

II.E. Role of Lipids Five lipids are associated with LHC II. Among them, digalactosyl diacyl glycerol (DGDG) and phosphatidyl glycerol (PG) appear to have special roles in the structure of the complex (see Chapter 1, Section VIII for a discussion of these lipids). It has been found that removal of DGDG by mild detergent treatment or anion exchange chromatography causes the complex to lose the ability to form two- or threedimensional crystals. The ability to crystallize can be completely restored, however, by readdition of pure DGDG at a ratio of 4 lipid molecules per polypeptide. Trimer formation appears to be a prerequisite for crystallization, as all attempts to form crystals of monomeric complexes have so far been unsuccessful. The lipid PG, on the other hand, binds more firmly than DGDG and cannot be removed by non-ionic detergent treatment, but only by delipidation with phospholipase or by proteolytic cleavage of the first 50 amino acids at the N-terminus. These treatments either break down the lipids or release them from the complex, in either case the result being the dissociation of the trimer into monomers. These results indicate that DGDG and PG have distinctly different and specific roles in the structure of the complex. PG is apparently involved in the formation of trimers and is likely located at the subunit interface, whereas DGDG presumably binds at the periphery of the trimers to maintain the structural integrity of the complex and to facilitate crystal formation. It is noted that at 3.4 Å resolution, none of these lipids has yet been located on the electron-density maps.

III. Regulatory Roles of LHC II in Thylakoids Besides the major role of harvesting light, LHC II also has two other important functions, one in the regulation of thylakoid membrane structure, i.e., grana stacking, and the other in a related phenomenon, the regulation of the distribution of excitation energy between the two photosystems.

IV. A. Regulation of Thylakoid Structure by LHC II Grana Stacking Apparently LHC II is essential for grana stacking, a notion supported by many lines of evidence. In developing chloroplasts, for example, the formation of grana stacks directly coincides with the appearance of LHC II. Furthermore, photosynthetic organisms and mutants which lack LHC II also lack grana stacks. In a barley mutant lacking Chl b and partially deficient in LHC II, the grana stacks are found to be less stable. Other supporting evidence is the reduced amount of grana stacking when LHC II is structurally altered by selective proteolytic cleavage of its surface-exposed segments.

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Grana stacking is an extraordinary phenomenon in structural biology, and it has intrigued photosynthesis researchers for decades. Grana stacking undoubtedly provides functional specialization to different regions of the membranes, particularly in regard to the physical separation of the two photosystems. It results in a lateral heterogeneity in the thylakoid membrane and as a consequence compartmentalization of the various photosynthetic complexes. Furthermore, it ultimately provides a means for increasing the overall efficiency in energy utilization by the two photosystems8,9.

IV. B. Regulation of Excitation-Energy Distribution by LHC II In oxygenic photosynthesis the linear electron transfer from to requires both photosystems II and I acting in series. Maximum efficiency for this linear electron transfer requires that the processes in both photosystems be balanced or, in other words, excitation of the two photosystems should occur at equal rates. During the late 1960s, evidence was obtained by Murata10 and by Bonaventura and Myers11 that an in vivo mechanism exists to ensure such a balanced excitation. This regulation was referred to as a “state 1-state 2 transition.” For instance, if PS II were receiving more excitation than PS I, either because of the available light intensity or its spectral distribution, the result would be accumulation of a pool of reduced plastoquinone, as the PS I-driven oxidation of would be rate-limiting. Under such a condition, however, the photosynthetic apparatus is apparently able to sense the situation and regulate the distribution of energy to achieve a balanced excitation of the two photosystems. In the late 1970s, Bennett12 found that the state transition is accompanied by a biochemical event. As reduced plastoquinone is accumulated when PS II is excited to a greater extent than PS I, light that overexcites PS II activates the membrane-bound kinase. The result is the phosphorylation of the surfaceexposed portion of LHC II, resulting in a partial destacking of the grana partitions and the migration of the LHC II from the grana region toward the stroma-exposed region. This migration reduces the excitation of the reaction centers of photosystem II and increases the proportion of the absorbed excitation energy that is transferred from the light-harvesting complexes to photosystem I. Conversely, when PS I is excited to a greater extent than PS II, becomes oxidized and kinase action is switched off. A membrane-bound phosphatase then acts to dephosphorylate the phosphorylated LHC II, and excitation of PS-II reaction centers is increased relative to that of PS I. Thus the distribution of excitation energy between the two photosystems can be regulated by a mechanism that is dependent on the redox state of the plastoquinone pool and involves the phosphorylation and dephosphorylation of LHC II. The above-mentioned changes have been supported by experiments carried out by Telfer, Allen, Barber and Bennett13. A controlled light regime was used with pea chloroplasts to cause a state transition and the amount of LHC II phosphorylated or dephosphorylated was then monitored by measuring the LHC II radio-labeled with 32P from ATP during the transition. In Fig. 7 (A), left, state 1 -to-state 2 transition was effected by first applying both (blue light, modulated at 90 Hz for the purpose of detecting fluorescence) and (unmodulated 710-nm light) and then after a few minutes turning off only . During the period when only is on, reduced plastoquinone is accumulated, as judged by the increase in the modulated fluorescence signal [not shown here, but see (B)], and the amount of radio32 P-labeled LHC II increased, as shown in Fig. 7 (A), left. The degree of phosphorylation was measured by the total radioactivity of the band on SDS-PAGE corresponding to the LHC-II. When only was applied initially and then was later superimposed, the state 2-to-state 1 transition occurs, and phosphorylated LHC II becomes dephosphorylated [see Fig. 7 (A), right]. These results provide clear support for the hypothesis that the transition to state 2 involves phosphorylation while the transition to state 1 involves dephosphorylation of LHC II. In other words, LHC II phosphorylation is controlled by the redox state of the quinone pool.

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An equally compelling piece of evidence that protein phosphorylation is controlled by the redox state of plastoquinone pool was provided by Horton, Allen, Black and Bennett14 using potentiometric titration of both fluorescence and LHC II phosphorylation, as shown in Fig. 7 (B). Details on the use of redox titration of PS-II fluorescence-yield changes to monitor the redox-state changes of the plastoquinone pool in PS II will be discussed in Chapter 17. The data points for fluorescence as well as for phosphorylation, as measured by the level of radioactive 32P-labeled LHC II, fit the Nernst equation [see footnote on p. 92] with an (at pH 7.8) of ~0 mV and an n-value of 2, reflecting a two-electron change for the electron carrier involved, presumably PQ. Thus crucial links are established between the redox state of plastoquinone, the activation of kinase for LHC II phosphorylation and changes in chlorophyll fluorescence.

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As it is known that the 25-kDa polypeptide of LHC II (o) is phosphorylated at a far higher rate than the 27-kDa polypeptide of the LHC II (i) complex, it is expected that LHC II(o) would be preferentially involved in phosphorylation. It is also known that phosphorylation targets the surface-exposed threonine residue (Thr6) near the N-terminus of the LHC II polypeptide. A proposed molecular mechanism responsible for regulation of excitation-energy distribution through phosphorylation and dephosphorylation of LHC (o) has been provided by Staehelin and Arntzen15, as shown in Figure 9, and also by Allen 16 . Starting with predominantly dephosphorylated illumination by favors turnover of PS II and reduction of plastoquinone pool which in turn activates kinase resulting in LHC II (o) phosphorylation. In the nonphosphorylated state, LHC II particles promote membrane stacking and therefore accumulate in the stacked membrane regions. Addition of phosphate group(s) to the exposed polypeptide segment adds negative charge to the surface and consequently produces electrostatic repulsion. As a result of the mutual repulsion between the negative charges on the phosphorylated LHC II (i.e., LHC IIP) on adjacent membranes, unstacking occurs and LHC II-P migrates out of the stacked region into the unstacked Such lateral migration of LHC II enables a redistribution of energy to re-establish a balanced excitation of the two reaction centers. This mechanism has received ample experimental support. Examination of freeze-fracture electron micrographs of nonphosphorylated, phosphorylated and dephosphorylated thylakoids created under appropriate experimental conditions revealed that ~20% of the mobile LHC II particles are involved in

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reversible migration between stacked and unstacked membrane regions. Additionally, fractionation of phosphorylated chloroplast membranes showed a tenfold higher relative phosphorylation in unstacked, stromal membranes than in stacked grana. When the viscosity of the medium for the thylakoid membrane is increased (with cholesteryl hemisuccinate), kinase activity is unaffected, but lateral migration of the mobile LHC II and the consequent state 1-to-state 2 transition are prevented.

For further reading: R1. DJ Simpson and J Knoetzel (1996) Light-harvesting complexes of plants and algae: Introduction, survey and nomenclature. In: DR Ort and CF Yocum (eds) Oxygenic Photosynthesis, 493-506. Kluwer R2. S Jansson (1994) The light-harvesting chlorophyll a/b-binding proteins. Biochim Biophys Acta 1184: 1-19 R3. W K hlbrandt (1994) Structure and function of the plant light-harvesting complex, LHC II. Current Opinion in Structural Biology. 4: 519-528 R4. LA Staehelin and GWM van der Staay (1996) Structure, composition, functional organization and dynamic properties of thylakoid membranes. In: DR Ort and CF Yocum (eds) Oxygenic Photosynthesis, 11-30. Kluwer R5. Bassi, F Rigoni and GM Giacometti (1990) Chlorophyll binding proteins with antenna function in higher plants and green algae. Photochem Photobiol 52: 1187-1206

References: 1.

2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.

S Jansson, E Pichersky, R Bassi, BR Green, M Ikeuchi, A Melis, DJ Simpson, M Spangfort, LA Staehelin and JP Thornber (1992) A nomenclature for the genes encoding the chlorophyll a/b-binding proteins of higher plants. Plant Mol Biol Reporter 10: 242-253 T Ogawa, F Obata and K Shibata (1966) Two pigment proteins in spinach chloroplasts. Biochim Biophys Acta 112: 223-234 JP Thornber, CA Smith and JL Bailey (1966) Partial characterization of two chlorophyll-protein complexes isolated from spinach-beet chloroplasts. Biochem J 100: 14-15 W K hlbrandt (1987) Three-dimensional crystals of the light-harvesting chlorophyll a/b-protein complex from pea chloroplasts. J Mol Biol 194: 757-767 W K hlbrandt and DN Wang (1991) Three dimensional structure of plant light-harvesting complex determined by electron crystallography. Nature 350: 130-134 R Henderson and PNT Unwin (1975) Three-dimensional model of purple membrane obtained by electron microscopy. Nature 257: 28-32 W K hlbrandt and DN Wang and Y Fujiyoshi (1994) Atomic model of plant light-harvesting complex by electron crystallography. Nature 367: 614-621 LA Staehelin and CJ Arntzen (1983) Regulation of chloroplast membrane function: Protein phosphorylation changes the spatial organization of membrane components. J Cell Biol 97: 1327-1337 H-W Trissl and C Wilhelm (1993) Why do thylakoid membranes from higher plants form grana stacks? Trends Biochem Sci 18: 415-419 N Murata (1969) Control of excitation transfer in photosynthesis. I. Light-induced change of chlorophyll a fluorescence in Porphyridium cruentum. Biochim Biophys Acta 172: 242-251 C Bonaventura and J Myers (1969) Fluorescence and oxygen evolution from Chlorella pyranoidosa. Biochim Biophys Acta 189: 366-383 J Bennett (1977) Phosphorylation of chloroplast membrane polypeptides. Nature 269: 344-346 A Telfer, JF Allen, J Barber and J Bennett (1983) Thylakoid membrane phosphorylation during state 1- state 2 transitions in osmotically shocked pea chloroplasts. Biochim Biophys Acta 722: 176-181 P Horton, JF Allen, MT Black and J Bennett (1981) Regulation of phosphorylation of chloroplast membrane polypeptides by the redox state of plastoquinone. FEBS Lett 125: 193-196 LA Staehelin and CJ Arntzen (1983) Regulation of chloroplast membrane function: Protein phosphorylation changes the spatial organization of membrane components. J Cell Biol 97: 1327-1337 JF Allen (1992) Protein phosphorylation in regulation of photosynthesis. Biochim Biophys Acta 1098: 275-335

Chapter 13 Role of Carotenoids in Photosynthesis I. The Light-Harvesting Role of Carotenoids II. Chlorophyll-Carotenoid-Protein Supramolecular Assemblies Xanthosomes A. Structure and Function of the Peridinin-Chlorophyll-Protein (PCP) Complex B. The Supramolecular Fucoxanthin-Chlorophyll-Protein Assembly (FCPA) 1. Spectral Properties 2. A Molecular Model III. Singlet-Singlet Energy-Transfer Dynamics in Light-Harvesting Carotenoids A. Low-Lying Excited Singlet States in Carotenoids B. Carotenoid Fluorescence C. Time-Resolved Spectroscopic Measurements of Carotenoid-to-Bacteriochlorophyll Energy Transfer in Bacterial Light-Harvesting Complexes IV. The Photoprotective Role of Carotenoids Triplet-Triplet Energy Transfer

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More than 600 naturally occurring Carotenoids are known at the present time. About 150 different carotenoids can be found in higher plants and such other photosynthetic organisms as algae and photosynthetic bacteria. Carotenoids as a class have exceedingly diverse functions in photosynthetic organisms but as depicted schematically in Fig. 1, they serve two major recognized functions in photosynthesis: one, as an accessory light-harvesting pigment and, the other, as a triplet quencher to provide protection against photooxidative damage. Increasing evidence also shows that carotenoids may also have a structural role (see Chapter 1). In their light-harvesting role, as depicted in Fig. 1 (A), a carotenoid in the antenna complex absorbs visible light energy that is not efficiently absorbed by chlorophyll, particularly, in the green, becoming transformed into the excited singlet state and transferring this excitation energy to either Chl or BChl molecules. This energy, originally absorbed by a carotenoid, is then rapidly transferred via chlorophylls to the reaction center to perform photochemistry. The efficiency of this singlet-singlet energy transfer [labeled “S-S” in Fig. 1 (A)] in native pigment complexes varies, depending on the structure and excitation energy of the carotenoid, as well as on its distance and orientation relative to the chlorophyll molecules. The efficiency of singlet-singlet energy transfer from carotenoid to chlorophyll is usually higher than 70%, and in some instances, such as excitation transfer in the B800-850 light-harvesting complex of sphaeroides [see Fig. 3 (C) below], it is close to 100%. Fig. 1 (B) illustrates the second major function of carotenoids, namely, photoprotection of chlorophyll, either by direct triplet-triplet energy transfer [labeled “T-T”] from to the carotenoid or by indirect transfer via the formation of singlet oxygen. The reaction steps for each of the two mechanisms are shown in Fig. 1, bottom, right. Further details on both the light harvesting by carotenoids and photoprotective role of carotenoids will be described in the following sections.

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A relationship between the role played by a carotenoid and its molecular configuration has been found recently. Yasushi examined the carotenoids acting as accessory pigments in light-harvesting complexes and those present in reaction centers where they act as triplet quenchers. A variety of spectroscopic techniques, including electronic absorption, Raman spectroscopy, and nuclear magnetic resonance, have all been used to establish the configuration of the carotenoids involved in a number of photosynthetic bacteria. Available evidence shows that an apparent natural selection of the molecular configuration prevails in accordance with the particular role. The carotenoids present in a reaction center (RC) usually assume the15-cis configuration, with some out-of-plane twist along the conjugated backbone, as illus-

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trated in the examples of sphaeroides and rubrum shown in Fig. 2. The cis-configuration of carotenoids in the bacterial reaction center has also been confirmed by X-ray crystallography. On the other hand, carotenoids in light-harvesting complexes of photosynthetic bacteria assume an alltrans configuration, either twisted (as in rubrum, Chromatium vinosum, and palustris) or planar (as in sphaeroides and capsulata). Note that chain twisting is not ascribed to the type of carotenoid, but rather to some specific interaction between the carotenoid and the protein environment and apparently affects the efficiency of singlet energy transfer. It is worth noting that Koyama’ s rule on carotenoid configuration in photosynthetic reaction centers first appeared to apply only to photosynthetic bacteria, since an earlier examination of the green-plant PS-II reaction-center complex led Fujiwara, Hayashi, Tasumi, Kanji, Koyama and to conclude that the assumes an all-trans configuration. However, the realization that the 15-cis form of is extremely unstable when excited by light to the triplet state, from which it can rapidly isomerize to the all-trans form, prompted Bialek-Bylka, Tomo, Satoh and Koyama2 to reexamine the PS-II reaction-center complex by using new procedures and new detection methods. Solvent extraction of was carried out in complete darkness at Spectrometric analysis of the extracted pigment by HPLC was also carried out at and conducted using an apparatus equipped with a two-dimensional diode-array detector for simultaneous detection and recording of electronic spectra for minimum exposure to light. With these precautions, the authors could spectroscopically identify 15-cisin the reaction center of photosystem II in spinach. Thus, the Koyama-rule correlating carotenoid configuration with its function now seems firmly established.

I. The Light-Harvesting Role of Carotenoids The peridinin-chlorophyll a -protein (PCP) complex in marine dinoflagellates is a well-known lightharvesting protein complex, whose structure and properties will be discussed in detail in Section II. Peridinin, an allenic xanthophyll, will be discussed in Section II. A. For the moment, we will discuss PCP only to illustrate the efficiency of energy transfer from carotenoid to chlorophyll in a light-harvesting complex. The light-harvesting role of carotenoids can be readily demonstrated by monitoring the fluorescence of chlorophyll when exciting, e.g., the PCP complex in a wavelength region where carotenoid is the principal absorber. In the absorption spectrum of PCP in Fig. 3 (A), top panel, the absorption maximum at 476 nm is due mainly to peridinin while the maximum at 435 nm is due mainly to chlorophyll a and that at 669 nm exclusively to chlorophyll a. Fluorescence excitation and emission spectra of PCP are shown in the left and right portions, respectively, of the bottom panel in Fig. 3 (A). The quantum-corrected excitation spectrum between 260 and 600 nm appears almost identical to the absorption spectrum, indicating efficient energy transfer from peridinin to chlorophyll. The uncorrected chlorophyll emission spectra in the right portion were elicited by excitation at 330 nm and 479 nm, as indicated. Separate fluorescence excitation measurements by Song, Koka, Prézelin and Haxo3 with PCPs from Glenodinium sp., Gonyaulax polyedra and Amphidinium rhyncocephaleum showed that energy transfer from peridinin to Chl a in all three algae was 100% efficient. In contrast, energy transfer between isolated chromophores of peridinin and Chl a together could not be observed in (dilute) solution. Also, denatured PCP showed no significant energy transfer, even though the peridinin chromophores might still have been attached to the protein. As a second example [Fig. 3 (B)], we will discuss the light-harvesting complexes of the brown alga Acrocarpiapaniculata (a seaweed). Two light-harvesting complexes were isolated by treatment with Triton X-100 followed by sucrose density-gradient centrifugation: a major orange-brown “B” fraction and a minor green “C” fraction, with the B-fraction containing ~60% of the total pigments and most of the carotenoid fucoxanthin.

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The absorption spectrum of fucoxanthin has peaks at 424, 444, and 470 nm in organic solvents, but is red-shifted in the pigment-protein complex. Fraction-B shows absorption peaks at 436 and 465 nm due to Chl a and Chl respectively, and major absorption bands in the 500-550 nm region due to fucoxanthin. The fluorescence excitation and emission spectra at 77 K for the (Fraction-B) and the (Fraction-C) are shown in Fig. 3 (B). Both excitation spectra were measured by monitoring the fluorescence at 680 nm arising from Chl a. The main features of the excitation spectra of the two complexes are rather similar and directly reflect the pigment composition of each. For example, the excitation spectrum of fraction C, which contains some fucoxanthin, shows a moderate amount of energy transfer in the 500-560 nm region, while the excitation spectrum for fraction-B apparently shows no contribution by violaxanthin. The excitation spectra also closely match the absorption spectra of the respective complexes (not shown) and clearly indicate that and fucoxanthin all transfer energy to the fluorescence emitter, Chl a. The third example is the light-harvesting complex B800-850 obtained from the wild-type sphaeroides 2.4.1. This complex contains three BChls and one carotenoid (spheroidene) per protein subunit. The fluorescence excitation was obtained by monitoring the emission at 850 nm. Fig. 3 (C) shows the absorption and fluorescence excitation spectra in the 400-620 nm region, with the two spectra normalized at 590 nm (marked with The excellent match of the absorption and excitation spectra indicates that photoexcited spheroidene transfers energy to bacteriochlorophyll with a high efficiency.

II. Chlorophyll-Carotenoid-Protein Supramolecular Assemblies – Xanthosomes For the efficient capture of low-intensity light, the light-harvesting complexes in certain species are present in the form of supramolecular assemblies consisting of intricate structures containing large number of pigment molecules. Examples of such supramolecular assemblies are the phycobilisomes in either cyanobacteria or red algae (Chapter 14) and the chlorosomes (Chapter 8) in green bacteria. It has been found recently that brown algae, which are usually found in deep waters where the available light intensity is especially low at physiologically important wavelengths, appear to possess such supramolecular, chlorophyll-carotenoid assemblies, the so-called “xanthosomes”4. Two unique xanthosomes will be discussed in this section. One is the peridinin-chlorophyll-protein (PCP) present in certain marine algae, the dinoflegellates. PCP is responsible for the color of the so-called “red tides.” Peridinin, a xanthophyll, and the principal pigment in the light-harvesting complex, augments light absorption by the organism in the blue-green region (470-550 nm), as light in this spectral region is little absorbed by chlorophylls. The other xanthosome is the “fucoxanthin-chlorophyll protein assembly” (FCPA) isolated rather recently by Tetzuya Katoh and colleagues5.

II. A. Structure and Function of the Peridinin-Chlorophyll-Protein (PCP) Complex Peridinin was isolated in 1890 by Franz Schütt6, who called it “peridineen” and even suggested that it was conjugated to a protein in vivo. In 1963, Bode and Hastings7 prepared an aqueous extract from Gonyaulax polyedra and attributed its red color to protein-bound peridinin. The molecular weight of this red protein was later reported by Haidak, Mathews and Sweeney8 to be ~38 kDa. A more highly purified sample was subsequently obtained by Haxo, Siegelman and their coworkers9, 10. Isoelectric focusing, however, revealed that there were actually various kinds of PCPs with closely related values of pI ranging from 6.5 to 7.5, the predominant species (90%) having a pI of 7.5. The molecular structure of peridinin itself was found by Harold Strain and coworkers11 in 1971 to be an asymmetric, tricyclic carotenoid, consisting of a cyclohexanol ring and an epoxycyclohexanol ring, connected by a polyene chain containing a furane ring:

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The peridinin-chlorophyll-protein complex can be easily released by a freeze-thaw cycle and then grinding the algal mass, after being refrozen. The purified PCP antenna complex from Amphidinium carterae has a molecular mass of~39 kDa, each subunit having 2 Chl-a molecules and 8 peridinins (previously reported as 9). On the other hand, the PCP complex isolated from either of the dinoflagellates Glenodinium sp., Gonylaulax polyedra, and Amphidinium rhynocephaleum consists of only 1 Chl a and 4 peridinin per protein subunit. Evidence for energy transfer in dinoflegellate PCP has already been presented in Fig. 3. In 1976, Song, Koka, Prézelin and Haxo3 and in 1977 Koka and Song12 measured the CD spectra of PCPs isolated from various dinoflegellates, with peridinin-Chl a-protein ratios of 4:1:1 or 8:2:1, and observed a splitting of the main absorption band of peridinin into two bands of approximately equal ellipticity but of opposite sign at~375 and~495 nm in both A. carterae and A. rhyncocephalem, as shown in Fig. 4 (A) [cf. the absorption and fluorescence excitation spectra of PCP from A. carterae in Fig. 3 (A)]. Note that the CD signal is completely lost when the PCP from Glenodinium sp. is denatured, as shown in Fig. 4 (B). The CD results suggest that four peridinin molecules form two dimers with a chlorophyll located between them. In each dimer in the native PCP the two peridinins are mutually orthogonal to one another. As previously noted, the peridinins are strongly coupled with chlorophyll allowing for efficient energy transfer. In addition, results from fluorescence-polarization measurements12 indicate that both components ofthe peridinin CD bands are oriented relative tothe axis ofchlorophyll a, as shown in the model in Fig. 4 (C).

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About twenty years later, in 1996, Hofmann, Wrench, Sharpies, Hiller, Werte and Diederichs12 crystallized the PCP complex from A. carterae and determined its three-dimensional structure by X-ray crystallography at 2.0 Å resolution. A stereo view ofthe PCP-monomer shown in Fig. 5 (A) [also see Color Plate 7] has a topology resembling a wide-open, right-handed “jelly roll,” with a short, loop forming the tip of the jelly roll. Such a jelly-roll fold had not previously been reported for an protein. The electron-density maps revealed the presence not only of water molecules but surprisingly also the presence of two digalactosyl diacyl glyceride (DGDG) lipid molecules, apparently forming an integral part of the PCP structure. The N- and C-terminal halves of the polypeptide form two almost identical domains related to each other by a twofold symmetry axis, each domain consisting of 8 helices of the polypeptide. Hofmann et al.13 likened the PCP monomer to a boat, with a bow, stern, deck and keel as indicated in the figure, the “keel” being formed by a 40 Å-long chain connecting the N- and C-terminal domains. The central hydrophobic cavity is occupied by the chromophores (2 Chl-a and 8 peridinins), like “cargos” inside the boat. Two large openings are found near the bow and the stern, between helices N6 and N8 and likewise between helices C6 and C8. Two smaller openings are also found near the deck, one between N2 and N7 and the other between C2 and C7, the first is occupied by the hydrophilic epoxycyclohexane ring of

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peridinin and the second by the head group of the lipids. The presence of two peridinin pairs surrounding a chlorophyll, as previously deduced from CD spectroscopy12, has now been confirmed by X-ray crystallography which also showed that the four peridinins are present in the all-trans configuration. Within each peridinin pair, the polyene chains are less than 4 Å apart at closest approach,. In contrast to the model based on CD results, however, the angle formed by the peridinins within each pair is and the furane rings in a pair are quite close to one another, the distance between their centers being 5.5 Å, vs. the 12 Å estimated previously12 for peridinins in each pair. As shown in the stereogram of Fig. 5 (B), the conjugated portion of each peridinin is close to the chlorophyll tetrapyrrole ring at a van der Waals distance (3.3 to 3.8 Å), a proximity which allows for efficient excitonic energy transfer from peridinin to chlorophyll. Within the PCP monomer, the two chlorophylls are 17.4 Å apart and Förster energy transfer should prevail. Like other light-harvesting protein complexes, such as the bacteriochlorophyll-a protein discussed in Chapter 8, the LHC II complex discussed in Chapter 12, and the phycobiliproteins to be presented in the next chapter, PCP monomers also form a trimer. The trimer 100-Å across and 40 Å thick is stabilized by hydrophobic interactions between the deck and stern helices and through hydrogen bonding of the peridinin epoxycyclohexane ring in the C-terminal domain. Within the trimer, the distances between chlorophylls of different subunits are in the 40-45 Å range, well within the range of dipole-dipole interactions required for Förster energy transfer. The precise location of the PCP complex within the membrane and how it transfers energy to the photosystems are not yet known, but circumstantial evidence suggests that PCP is located in the intrathylakoid space. Hofmann et al.13 have suggested that PCP is probably linked to the reaction center by way of a membrane-bound LHC. In 1990, Mimuro, Tamai, Ishimaru and Yamazaki14 examined transfer of excitation energy in the marine dinoflegellate Protogonyaulax tamarensis by means of both steady-state and time-resolved fluorescence spectroscopy. At ambient temperature, one dominant fluorescence band was found at 684 nm (F684) with a minor band at 673 nm (F673), irrespective of whether peridinin, Chl a or Chl c was preferentially excited. At 77 K, three major fluorescence bands F670, F683, and F689 originating from Chl a in photosystem II were resolved. The rise and decay kinetics of these low-temperature bands indicated that energy absorbed by peridinin is transferred first to F670 in < 22ps, then to F683 and finally to F689. The strongest emission band, F689, in the dinoflegellate was considered to be the equivalent of F695, the characteristic band of the core antenna of green-plant PS II. These time-resolved fluorescence kinetics led Mimuro et al.14 to suggest the following energy-transfer sequence: These authors also found evidence for energy transfer to photosystem I via spill-over, though not through F689. No evidence was found for involvement of Chl c in energy transfer from peridinin to Chi a. Hofmann et al.13 also suggested that trimers of PCP and (PS-II)LHC might be stacked over one another, thus allowing a highly efficient, Förster energy transfer from PCP to (PS-II) LHC.

II.B. The Supramolecular Fucoxanthin-Chlorophyll-Protein Assembly (FCPA) Until recently, the chlorophyll-carotenoid light-harvesting complexes were not known to form supramolecular complexes comparable to the phycobilisomes or chlorosomes. About 10 years ago, however, Tetzuya Katoh and coworkers5, 15, 16 isolated an orange-brown, supramolecular, light-harvesting complex containing chlorophylls a and c and fucoxanthin from the brown alga Dictyota dichotoma5 using the detergent decanoyl glucoside, and subsequently also from another brown alga Petalonia fascia15, both complexes similar in several respects to the supramolecular phycobilisomes and chlorosomes. The authors called this new, 520-kDa complex “fucoxanthin-chlorophyll-protein assembly,” abbreviated as FCPA.

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II. B. 1. Spectral Properties The FCPA complex is made up of seven protein subunits, each with a molecular weight of 74 kDa and each containing 13 molecules of Chl a, 3 Chl c, 10 fucoxanthin and 1 violaxanthin. The pigment content of the FCPA complex is unusually high, with a pigment-to-protein weight ratio of 0.37. The intact, orange-brown FCPA complex is readily dissociated into its seven protein subunits by the action of detergents such as LDAO or Triton X100 at very low concentrations, even as low as 0.005%. The extreme frailty of this supramolecular complex may explain why it had not been isolated earlier. The absorption spectra of the intact FCPA and its second-derivative spectra are shown for room temperature in Fig. 6 (A), top and at 77 K in Fig. 6 (A), bottom. Note that the derivative spectra are shown not in the conventional manner but up-side-down (as so that the positive peak in the secondderivative trace corresponds to a normal absorption peak and a negative-going trace corresponds to a shoulder. The major peaks at room temperature belonging to Chl a are located at 439 and 671 nm; Chl c has a peak at 636 nm and a shoulder at 459 nm. The presence of fucoxanthin is represented by small shoulders at 515 and 545 nm, indicating that it is present in its long-wavelength forms. These bands are clearer in both low-temperature and second-derivative spectra. In the low-temperature spectra, both the blue and red bands of Chl c are split. Many more minor bands are resolved for fucoxanthin in the derivative spectra, suggesting a high degree of “heterogeneity.”

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The absorption spectra of the intact FCPA and dissociated FCPA subunits are shown in Fig. 6 (B), top and the difference spectrum obtained by subtracting intact FCPA spectrum from Triton-treated spectrum in Fig. 6 (B), bottom. In the red region, the major change is a blue shift ofthe Chl a absorption upon Triton treatment and The other major change is a blue shift in the fucoxanthin absorption and although the latter is partially offset by the Chl a absorption decrease. The difference in the fluorescence spectra between intact and dissociated FCPA is rather dramatic, as can be seen in Fig. 7. The main emission band at 677 nm shown in Fig. 7 (A) was excited at 440, 460 and 515 nm, wavelengths absorbed predominantly by Chl a, Chl c and fucoxanthin, respectively. The emission spectra indicate that both Chl c and fucoxanthin transfer excitation energy to Chl a with high efficiency. The emission spectrum excited by 460 nm shows a weak emission at 640 nm, originating from Chl c. However, the 640-nm emission is not seen when the sample was excited at 515 nm, which suggests that fucoxanthin transfers energy directly to Chl a without the mediation of Chl c. In the dissociated FCPA, excitation at 440, 460 and 515 nm evokes quite different emission patterns [Fig. 7 (B)]. The 460-nm light, preferentially absorbed by Chl c, produces an increase in the emission at 640 nm and much less emission at 670 nm. Similarly, excitation by 515-nm light absorbed primarily by fucoxanthin also produced very little 677-nm emission but no emission at all at 640 nm. These results suggest that the coupling between Chl c and Chl a and that between fucoxanthin and Chl a are both disrupted and they also confirm that transfer of energy from fucoxanthin to Chl a does not go through Chl c.

II.B.2. A Molecular Model An electron microscope examination of the intact FCPA isolated from Petalonia fascia showed the supramolecular assemblies to be uniform in size and shape. They were found to be discoidal, 11.2 nm in diameter and 10.2 nm in height, and with a cavity in the center. Long-term storage of FCPA resulted in the

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formation of an amorphous material and in the loss of fine structure in its electron micrograph. Since FCPA is hydrophobic, it is presumed that the complex is partly immersed in the thylakoid membrane, rather than being attached to the membrane surface as is the case for the phycobilisomes or chlorosomes. Topological details of the native supramolecular complex remain to be investigated. With additional spectroscopic studies Mamoru Mimuro and coworkers16 were able to construct a molecular model for the FCPA complex. As already noted, spectrally heterogeneous forms of chlorophylls a and c and fucoxanthin are evident in the second-derivative and circular-dichroism spectra. Deconvolution of the absorption spectra revealed two chlorophyll-a forms, and Among the 13 Chl a molecules found in intact FCPA, 10 are and 3 are Two forms of Chl c were also found, a short wavelength-absorbing designated as and a long wavelength-absorbing designated as Further analysis of the absorption spectra indicated that there are two molecules of and one per subunit. Functionally active fucoxanthin absorbing at 515 and 545 nm and inactive fucoxanthin absorbing at 428, 455 and 488 nm were also found. Significantly, there was no indication of a strong interaction between molecules of any given pigment type. Molecular heterogeneity, inferred on the basis of spectral heterogeneity, and the absence of such interaction led Mimuro et al.16 to conclude that energy transfer occurs by direct coupling in one-to-one manner in donor-acceptor pigment pairs of the following four types:

The numbers inside the parentheses indicate the number of each type of pigment pair per subunit, as suggested by the mutual orientation of dipole moments of the respective pigment molecules calculated from the results of fluorescence polarization experiments. Three Chl-c molecules are separately paired with three as shown in Fig. 8. Two of the three are paired with one fucoxanthin each. Since both intact and dissociated FCPA have fluorescence emission maxima at 677 nm, it is clear that even though coupling between Chl c and and between fucoxanthin and are disrupted in dissociated FCPA, two molecules are transformed into and energy transfer among molecules and between and the terminal emitter (written as ) can still occur. In this model, the function of one remaining fucoxanthin and one violaxanthin has not yet been determined (see Fig. 8). By comparing the fluorescence excitation spectrum with the absorption spectrum of FCPA, it is seen that a decrease of 15-25% in transfer efficiency appears in the 480 nm-region, which would be consistent with an absence of coupling by two out of the 11 carotenoid molecules.

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III. Singlet-Singlet Energy Transfer Dynamics involving Carotenoids in Light-Harvesting Although energy transfer from carotenoids to chlorophylls in photosynthetic organisms can be readily demonstrated, as already seen in Fig. 3 of Section I, there has been a lack of understanding of the molecular mechanism by which the transfer takes place. First of all, isolated carotenoid molecules have been considered for some time to be virtually nonfluorescent and, thus, the Förster mechanism of energy transfer via dipole-dipole interaction, which requires not only an extensive overlap between the fluorescence spectrum ofthe energy donor and the absorption spectrum ofthe energy acceptor but also a high quantum yield of fluorescence for the isolated donor, is therefore not expected to be effective in the case of carotenoid-to-chlorophyll energy transfer.

III.A. Low-Lying Excited Singlet States in Carotenoids As a guide to understanding energy transfer involving carotenoids, we present an overview of the electronic states of carotenoids in relation to those of the polyenes, with the help of the energy diagrams of carotenoid, Chl a and BChl a in Fig. 9. The two excited states and of carotenoid are also denoted as and states, the customary electronic-state notations for states of different symmetry in linear polyenes. the ground state, is denoted as the state. The state now designated as had long been thought to be the lowest excited singlet state, as the state now designated as had not been observed in absorption or emission due to the fact that a transition is dipole forbidden, at least by a single photon process, while a transition is allowed, and in fact it accounts for the intense absorption band of carotenoids in the visible region. The existence of the state was shown by Thrash, Fang and Leroi 17 who measured the Raman excitation profile of in cyclohexane in the pre-resonance region, i.e., in the 16900-19000 region just below the region for the allowed transition, and found features that could be interpreted as evidence for a low-lying excited state 3470 below the state, i.e., with its origin near 17230 if the level is taken to be 20700 (483 nm). Since the fluorescent spectrum of would have a better spectral overlap with the long-wavelength Chl a absorption bands in the light-harvesting complex, it was interpreted by these authors18 as a possible energy donor transition in a Forster-type excitation transfer from carotenoid to antenna chlorophyll. But since the quantum yield ofthe donor emission an important term in the Förster mechanism, would be extremely small for this forbidden, or nearly forbidden, transition, Naqvi 19 suggested that energy transfer from excited to chlorophyll probably takes place by way of the “electron-exchange interaction” mechanism of Dexter20. Dexter’s electron-exchange interaction requires the electron in the highest occupied orbital of the excited donor and an electron acceptor to be in close contact i.e., form a “collision complex” so that exchange resonance interaction can occur via an overlap of their electron orbitals. This then allows the excited electron to move from to A while an electron in the highest occupied orbital of the ground-state of the acceptor, A, moves to D. It may be noted that exchange-resonance interaction requires close physical contact, i.e., overlap of wave functions, between the interacting partners, while the Förster energy transfer can operate over much longer distances (50-100 Å), by “through space” interactions. Furthermore, the electron-exchange mediated transfer is independent of the fluorescence quantum yield, The low quantum yield of fluorescence in carotenoids is due to an extremely rapid, non-radiative internal conversion followed by the fluorescence decay process The lifetimes of excited and states are of the order of 0.1 ps and 10 ps, respectively. The lifetime can

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be determined either directly by femtosecond spectroscopyR6, or indirectly by phase fluorometry. It is clear that the very short, non-radiative relaxation time makes it a more competitive deactivation process for the state, whereas the lower energy, state decays to the ground state in 10 ps, i. e., ~ 100 times longer, making carotenoid a more favorable candidate as the donor responsible for the light-harvesting role.

III. B. Carotenoid Fluorescence In the meantime, fluorescence excitation and emission spectra for isolated rhodopin, and spheroidene in were reported by Tomas Gillbro and Richard Cogdell21 in 1989. Their results for in Fig. 10 (A) show a very close match between the excitation and the absorption spectra, as well as extensive overlap between the excitation and emission spectra. The emission spectrum of which shows a small Stokes shift, is almost a mirror image of the absorption spectrum and apparently originates from the excited singlet state populated directly by light absorption, and not from a lower, indirectly populated state.. The quantum yield of fluorescence is notably very low, only for carotene and 3x10–5 for spheroidene. The fluorescence excitation and emission spectra of fucoxanthin were subsequently measured in 1991 both by Katoh, Nagashima and Mimuro 22 and by Shreve, Trautman, Owens and Albrecht23. The absorption and fluorescence spectra of fucoxanthin (isolated from brown algae) in are shown in Fig. 10 (B). The excitation spectrum (not shown here) for fucoxanthin fluorescence is almost identical to its absorption spectrum, with peaks located at 448,476 and 505 nm. The fluorescence spectrum excited by 476 nm light reaches a maximum at ~750 nm, a wavelength much longer than expected if the emission originated from the excited state The quantum yield of fucoxanthin fluorescence was estimated to be about 5x10–5, comparable to the yields for other carotenoids and therefore this Stakes-shifted emission most likely originated from the low-lying, excited singlet state

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Whether carotenoid fluorescence originates from the transition appears to depend on the number of conjugated double bonds, i.e., the chain length, and possibly other molecular-structural factors. For instance, the fluorescence from both and spheroidene, each of which has 10 conjugated double bonds, originates from the transition, while that of fucoxanthin, which has 8 conjugated double bonds, displays the Stokes-shifted emission. DeCoster, Christensen, Gebhard, Lugtenburg, Farhoosh and Frank24 performed a series of elegant experiments in which they examined the fluorescence spectra of four, all-trans spheroidenes containing 7 to 10 conjugated double bonds, and found that as the extent conjugation increases, there was a systematic crossover from emission to emission, i.e., from The fluorescence emission, excitation and absorption spectra of the four spheroidenes measured at room temperature are shown in Fig. 11. With more than 8 conjugated double bonds, fluorescence is dominated by emission. Starting with spheroidene with 10 conjugated double bonds, the fluorescence spectrum is almost a mirror image of the absorption. There is no evidence for the longer wavelength emission. The fluorescence spectrum of 3,4-dihydrospheroidene, which contains 9 conjugated double bonds, is similar to spheroidene, or but shows a very weak fluorescence in the 700-800 nm region. In the less conjugated carotenoid, 3,4,5,6-tetrahydrospheroidene, excitation gives dual fluorescence bands of comparable and intensity originating from both emissions. In 3,4,7,8-tetrahydrospheroidene, which contains only 7 conjugated double bonds, the fluorescence consists almost exclusively of the emission. The crossover in fluorescence from to Stokes-shifted emission is best explained by considering whether deactivation of the state takes the pathway of fluorescence or internal conversion. With increasing conjugation, experiment has shown that the energy of both and states decreases column in Fig. 11], while the gap between the two states, i.e., increases. This widening of the gap decreases the probability of the internal conversion process and, by

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default, increases the probability of fluorescence from the state. Thus, we see the general phenomenon of high-conjugation carotenoids being associated with the or emission, while low-conjugation carotenoids are associated with emission from the or state. Extrapolation of the transition energy indicates that the energy of the state of longer-chain carotenoids have considerably lower energy than previously thought, perhaps as low as ~14,000 for However, such a low estimated value for the -state energy of would exclude it from being the energy donor to the state of the chlorophyll acceptor (see Fig. 9).

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III.C. Time-resolved Spectroscopic Measurements of Carotenoid-to-Bacteriochlorophyll Energy Transfer in Bacterial Light-Harvesting Complexes Energy transfer from carotenoid in the B800-850 light-harvesting complex of purple photosynthetic bacteria sphaeroides is extremely efficient, approaching 100%. Time-resolved studies on energy transfer from carotenoid to BChl was first reported by Wasielewski, Tiede and Frank 25 for the B800-850 complex of acidophila. The authors used a 510 nm, 4-ps pulse to pump the absorption band of the carotenoid, rhodopin. The bleaching ofthe carotenoid ground state, i.e., at 480 nm occurs virtually instantaneously while the recovery occurs with a time constant of Within experimental error, the bleaching of BChl at 860 nm, i.e., its conversion into its first excited singlet state, occurred with the same time constant, namely The essentially identical times for the two processes are consistent with direct energy transfer from carotenoid to BChl in the antenna complex. One of the earliest femtosecond, time-resolved absorption spectroscopic studies was carried out by Shreve, Trautman, Frank, Owens and Albrecht26 in an investigation ofenergy transfer from the carotenoid spheroidene, in the B800-850 light-harvesting complex of sphaeroides. They used ~200 fs-duration pulses at 480,510 and 800 nm to excite the B800-850 antenna complex and found a decay time of ps for the B800 excited state, i.e., Since the quantum yield for B800-to-B850 energy transfer is known to be nearly unity, the decay of B800* is presumably due almost entirely to energy transfer to B850, i.e., Transient absorption changes of spheroidene in cyclohexane at 540-nm induced by 480-nm pulses showed the ground-state recovery, at room temperature to be 9.1 ps. However, similar scans made with a higher time resolution showed an initial bleaching (attributable to the transition)which rapidly decayed in 0.34 ps to a longer-lived state. The 0.34-ps lifetime of this state is consistent with the lifetime indirectly estimated from the quantum yield of fluorescence, On the basis ofthe dynamics ofthe electronic states of spheroidene in vitro, the authors suggested the possibility that energy transfer to B800-850 may occur from both the states of spheroidene. Under this assumption, the 800-nm transient absorption changes were examined by exciting the complex in the carotenoid spectral region. The results ofthese experiments are brought together in the kinetic model in Fig. 12. The transfer times were determined to be 1.7 ps and 3.8ps, respectively. In this two-channel model, the initially formed state is seen to decay by two routes. One is internal conversion to the state, and the other is energy transfer to BChl, i.e., B800. Since the internalconversion takes only 340f s , spheroidene must transfer energy from the state to B850 in ~300-400 f s . To account for the overall high efficiency of energy transfer, the remainder of the energy is assumed to be transferred to B800 from the state of spheroidene in competition with the internal conversion process As the latter process is much slower than the internal conversion, even a relatively slow energy transfer from can still be the dominant energy-transfer process.

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IV. The Photoprotective Role of Carotenoids – Triplet-Triplet Energy Transfer Excited singlet chlorophyll or bacteriochlorophyll can form their respective triplets by intersystem crossing. The otherwise slow decay of is accelerated by energy transfer (“quenching”) to triplet oxygen to form excited singlet oxygen a highly reactive species toxic to many cell constituents such as lipids, proteins, etc. Thus the second major property of Carotenoids with regard to photosynthesis is its ability to quench triplet (B)Chl at a much higher rate than by (triplet) oxygen, thus preventing formation of the very reactive singlet oxygen. Carotenoids can also quench, or scavenge, singlet oxygen once formed and thus provide additional protection. The pathways ofthese photoprotective functions have already been shown in Fig. 1. A more detailed scheme for the bacterial case is presented in Fig. 13, where Q is initially reduced. Photoexcitation of the bacterial reaction center promotes the primary donor into the first excited singlet state, which then transfers an electron to the primary electron acceptor, bacteriopheophytin, [step (a)]. Since the electron acceptor quinone, of the reaction center is chemically reduced (or absent), forward electron transfer is blocked. In this situation, the oxidized primary donor and the reduced recombine in ~10 ns, some of the reaction centers returning directly to the ground state, but many first forming the triplet state of the (re-reduced) primary electron donor [step (c)] via the triplet-state of the radical pair [step b]. The primary-donor triplet has a natural lifetime of ~50 ms but can be quenched by a carotenoid [step (d)]; it may also react with oxygen to produce singlet oxygen, although at a much slower rate than reaction with carotenoid [cf. Fig. 1 ]. The carotenoid triplet eventually reverts to the ground state with the release of heat [step (e)]. Usually, the direct quenching of the primary-donor triplet by carotenoid occurs two to three orders of magnitude faster than by oxygen, with an efficiency approaching 100% at room temperature. The quenching efficiency decreases as the temperature is lowered, decreasing by about 50% when the temperature has reached ~35-45 K, with almost no carotenoid triplet state formed at ~10 K. In the carotenoidless mutant R26 of sphaeroides in the absence of oxygen, the triplet decays to the ground state with a natural lifetime of ~50 ms. X-ray crystallographic studies have established the location of the carotenoid in the reaction centers of viridis [cf. Fig. 1 (B)] and two wild-type strains of sphaeroides. The lone carotenoid molecule (1,2-dihydroneurosporene in viridis, spheroidene in sphaeroides) is located on the M-subunit side,

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with the monomeric lying between it and the primary donor and ~3 Å from each (cf. Fig. 13). Because of the intermediate position of between, and its proximity to, the primary donor and the carotenoid, there has been a temptation to suggest that might serve as a conduit in the transfer of triplet energy. This possibility was examined experimentally by Harry Frank and Carol Violette27. Fig. 14 illustrates the treatment of a reaction-center preparation ofthe carotenoidless mutant sphaeroides with sodium borohydride to remove the and subsequent reconstitution with spheroidene. Examination of the reconstituted reaction center by absorption spectroscopy and circular dichroism demonstrated that the environment of the spheroidene incorporated into the reaction center was the same as in native sphaeroides. Spheroidene is apparently bound to the same site whether the monomeric BChl is present or not. EPR and transient optical absorption measurements showed, however, that triplet energy transfer from the primary donor to the carotenoid was inhibited in the borohydride-treated reaction center reconstituted with spheroidene, but lacking These observations provide direct evidence for the involvement of the monomeric BChl molecule in the triplet energy transfer pathway. Thus, the symmetrical bacterial reaction-center structure serves two functions in the early events of photosynthesis: normal electron transport on the L-, or A-, subunit side and triplet energy transfer on the M-, or B-, subunit side. The reaction center thus provides not only an effective means for achieving stable charge separation but also a mechanism for protecting the reaction-center complex from photodamage.

The understanding of the dynamics of triplet energy transfer has been greatly facilitated by the work of Larry Takiff and Steven Boxer29,30 who determined the first accurate values of triplet-state energies of the monomeric BChl a and the donor in the reaction centers of two photosynthetic bacteria from measurement ofphosphorescence intensities. The typical phosphorescence intensity ofbacteriochlorins, however, is exceedingly low: while the fluorescence quantum yield of BChl a is an easily measurable 0.21, the phosphorescence quantum yield has been found to be only as estimated from the intensities ofthe emission bands integrated over both time and wavelength. Measurements by these workers were made possible by a newly developed, ultrasensitive, germanium photodiode detector cooled by liquid nitrogen. Typical spectra of absorption, fluorescence and phosphorescence of BChl a in vitro are shown in Fig. 15 (A). The phosphorescence spectrum of the primary electron donor is illustrated for Rb. sphaeroides in Fig. 15 (B). Phosphorescence from the Rb. sphaeroides reaction center complex with the quinones removed was detected at 20 K on the long-wavelength side of the fluorescence and displayed an emission band with a maximum at 1318 nm (or 7590 and a half bandwidth of 240 The ratio of the integrated intensities of this 1318-nm luminescence to that of the fluorescence was, as expected, quite small, only

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Interestingly, as seen in Fig. 15 (B), phosphorescence was not observed with the reaction-center complex that still contained the quinone(s). The triplet-state energies of the BChl a and BChl b derived from the phosphorescence spectra are summarized for chromophores of Rb. sphaeroides and Rp. viridis at the appropriate levels in the diagram in Fig. 15 (C), together with those for the respective carotenoids, spheroidene (in Rb. sphaeroides) and 1,2-dihydroneurosporene (in Rp. viridis). Singlet oxygen is also shown.

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The triplet-state energy values determined by Takiff and Boxer account for the photoprotective function of carotenoids, based on their ability to quench the primary-donor triplets and thus prevent them from quenching the state of oxygen. It is known that the primary-donor pair in the bacterial reaction center is ~10 Å from the carotenoid, making it too far away for fast triplet-energy transfer. As Frank and Violette27 have shown, the monomeric BChl located between the primary donor and the carotenoid appears to mediate the formation of carotenoid triplet. The BChl-a triplet lies above the primary-donor triplet level, i.e., within of it at room temperature. Takiff and Boxer thus suggest that the longdistance transfer of triplet energy from the primary donor to the carotenoid occurs via a thermally-assisted transfer to the monomeric BChl first and then to the carotenoid. This proposal is also consistent with the fact that carotenoid triplet yield decreases sharply with decreasing temperatures, with the eventual elimination of triplet transfer all together at 10 K in the Rb. sphaeroides reaction centers. By contrast, the triplet level of the monomeric BChl b in Rp. viridis is more than above the primary-donor triplet making such a mechanism energetically unfavorable and thus triplet energy transfer from the primary donor to the carotenoid is prevented at all temperatures up to the room temperature.

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For further reading: R1. HA Frank, AD Young, G Britton and RJ Cogdell [eds] (1999) The Photochemistry of Carotenoids. Kluwer R2. A Young and G Britton [eds] (1993) Carotenoids in Photosynthesis. Chapman and Hall R3. Y Koyama, M Kuki, PO Andersson and T Gillbro (1996) Singlet excited states and the light-harvesting function of Carotenoids in bacterial photosynthesis. Photochem Photobiol 63: 243-256 R4. Koyama (1991) Structures and functions of carotenoids in photosynthetic systems. J Photochem Photobiol, B: Biol, 9: 265-280 R5. HA Frank and RJ Cogdell (1996) Carotenoids in photosynthesis. Photochem Photobiol 63: 257-264 R6. D Siefermann-Harms (1985) Carotenoids in photosynthesis. I. Location in photosynthetic membrane and lightharvesting function. Biochim Biophys Acta 811: 325-355

References: 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

15.

H Fujiwara, H Hayashi, M Tasumi, M Kanji, Y Koyama and (Ki) Satoh (1987) Structural studies on a photosystem II reaction center complex consisting of D-1 and D-2 polypeptides and cytochrome b-559 by resonance Raman spectroscopy and high-performance liquid chromatography. Chem Lett 10: 2005-2008 GE Bialek-Bylka, T Tomo, (Ki) Satoh and Y Koyama (1995) 15-cis-carotene found in the reaction center of spinach photosystem II. FEBS Lett 363: 137-140 P-S Song, P Koka, BB Prezelin and FT Haxo (1976) Molecular topology of the photosynthetic light-harvesting pigment complex, Peridinin-chlorophyll a-protein, from marine dinoflegellates. Biochemistry 15: 4422-2247 T Katoh, A Tanaka and M Mimuro (1993) Xanthosomes: Supramolecular assemblies of xanthophyll-chlorophyll a/c protein complexes. Methods in Enzymology 214: 402-412 T Katoh, M Mimuro and S Takaichi (1989) Light-harvesting particles isolated from a brown alga, Dictyota dichotoma. A supramolecular assembly of fucoxanthin-chlorophyll-protein complexes. Biochim Biophys Acta 976: 233-240 F Sch tt (1890) Ueber Peridineenfarbstoffe. Ber deutsch bot Ges 8: 9-32 VC Bode and JW Hastings (1963) The purification and properties of the bioluminescent system in Gonyaulax polyedra. Arch Biochem Biophys 103: 488-499 DJ Haidak, CK Mathews and BM Sweeney (1966) Pigment protein complex from Gonyaulax. Science 152: 212-213 FT Haxo, JH Kycia, GF Somers, A Bennett and HW Siegelman (1976) Peridinin-chlorophyll a protein of the dinoflegellate Amphidinium carterae (Plymouth 450). Plant Physiol 57: 297-303. HW Siegelman, JH Kycia and FT Haxo (1976) Peridinin-chlorophyll a-proteins ofdinoflegellate algae. Brookhaven Symp Biol 28: 162-169 HH Strain, WA Svec, K Aitzetm ller, MC Grandolfo, JJ Katz, H Kjłsen, S Norgard, S Liaaen-Jensen, FT Haxo, P Wegfahrt and H Rapoport (1971) The structure of peridinin, the characteristic dinoflegellate carotenoid. Am Chem Soc 93: 1823-1825 P Koka and P-S Song (1977) The chromophore topography and binding environment of peridininchlorophyllprotein complexes from marine dinoflegellate algae. Biochim Biophys Acta 495: 220-231 E Hofmann, PM Wrench, FP Sharples, RG Hiller, W Werte and K Diederichs (1996) Structural basis of lightharvesting by carotenoids: Peridinin-chlorophyll-protein from Amphidinium carterae. Science 272: 1788-1791 M Mimuro, A Tamai, T Ishimaru and I Yamazaki (1990) Characteristic fluorescence components in photosynthetic pigment system of a marine dinoflegellate, Protogonyaulax tamarensis, and excitation energy flow among them. Studies by means of steady-state and time resolved fluorescence spectroscopy. Biochim Biophys Acta 1016: 280-287 T Katoh and T Ehara (1990) Supramolecular assembly of fucoxanthin-chlorophyll-protein complexes isolated from a brown alga, Petalonia fascia. Electron microscopic studies. Plant Cell Phyiol 31: 439-447

250 16. M Mimuro, T Katoh and H Kawai (1990) Spatial arrangement of pigments and their interaction in the fucoxanthin-chlorophyll a/c protein assembly (FCPA) isolated from the brown alga Dictyota dichotoma. Analysis by means of polarized spectroscopy. Biochim. Biophys Acta 1015: 455. 17. RJ Thrash, HLB Fang and GE Leroi (1977) The Raman excitation profile spectrum of in the preresonance region: Evidence for a low-lying singlet state. J Chem Phys 67: 5930-5933 18. RJ Thrash, HLB Fang and GE Leroi (1979) On the role of forbidden low-lying excited states of light-harvesting carotenoids in energy transfer in photosynthesis. Photochem Photobiol 29: 1049-1050 19. KR Naqvi (1980) The mechanism of singlet-singlet excitation energy transfer from carotenoids to chlorophyll. Photochem Photobiol 31: 523-524 20. DL Dexter (1953) A theory of sensitized luminescence in solids. J Chem Phys 21: 836-860 21. T Gillbro and RJ Cogdell (1989) Carotenoid fluorescence. Chem Phys Lett 158: 312-316 22. T Katoh, U Nagashima and M Mimuro (1991) Fluorescence properties of the allenic carotenoid fucoxanthin: Implication for energy transfer in photosynthetic pigment systems. Photosynthesis Res 27: 221-226 23. AP Shreve, JK Trautman, TG Owens and AC Albrecht (1991) A femtosecond study of electronic state dynamics of fucoxanthin and implication for photosynthetic carotenoid-to-chlorophyll energy transfer mechanisms. Chem Phys 154: 171-178 24. B DeCoster, RL Christensen, R Gebhard, J Lutgenburg, R Farhoosh and HA Frank (1992) Low-lying electronic states of carotenoids. Biochim Biophys Acta 1102: 107-114 25. MR Wasielewski, DM Tiede and HA Frank (1986) Ultrafast electron and energy transfer in reaction centre and antenna proteins from photosynthetic bacteria. In: GR Flemming and AE Siegman (eds) Ultrafast Phenomena,Vol 5: 388-392. Springer Verlag 25. AP Shreve, JK Trautman, HA Frank, TG Owens and AC Albrecht (1991) Femtosecond energy-transfer processes in the B800-850 light-harvesting complex of Rhodobacter sphaeroides 2.4.1. Biochim Biophys Acta 1058: 280-288 27. HA Frank and CA Violette (1989) Monomeric bacteriochlorophyll is required for the triplet energy transfer between the primary donor and the carotenoid in photosynthetic bacterial reaction centers. Biochim Biophys Acta 976: 222-232 28. SL Ditson, RC Davis and RM Pearlstein (1984) Relative enrichment of P-870 in photosynthetic reaction centres treated with sodium borohydride. Biochim Biophys Acta 766: 623-629 29. LTakiff and SG Boxer (1987) Phosphorescence from the primary electron donor in Rhodobacter sphaeroides and Rhodopseudomonas viridis reaction centers. Biochim Biophys Acta 932: 325-334 30. L Takiff and SG Boxer (1987) Phosphorescence spectra of bacteriochlorophylls. J Am Chem Soc 110: 4254426

Chapter 14 Phycobiliproteins and Phycobilisomes I.

Phycobiliproteins A. Phycobilin Pigments B. Structure and Spectral Properties of Phycobiliproteins C. Crystal Structure of Phycobiliprotein Subunits II. Phycobilisomes A. The Linker Polypeptides B. Spatial Structure of Phycobilisomes in Anabaena variabilis C. Spectral Properties of Phycobilisomes D. Energy Transfer between Chromophores in Anabaena variabilis Phycobilisomes

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Light-harvesting complexes that drive photosynthesis are present in diverse forms. The Chl a/b-binding proteins discussed in Chapter 12 are proteins integral to the thylakoid membrane (see Fig. 1, left). However, in the prokaryotic cyanobacteria (blue-green algae) and eukaryotic red algae, light harvesting is carried out in the main by supramolecular assemblies called “phycobilisomes” (PBS) which are attached to the stromal surface of the photosystem-II core in the thylakoid membrane (see Fig. 1, right). Each phycobilisome consists of a group of brilliantly colored “phycobiliproteins” (PBP), each of which in turn contains covalently bonded pigments or chromophores called“phycobilins” (PB). The primary function of this remarkable light-harvesting apparatus is to allow the organism to survive in weak light conditions. As green and yellow light is transmitted unattenuated through great depths of water where the organisms reside, phycobilisomes possess the particular ability to absorb photons in these spectral regions where light is only weakly absorbed by chlorophyll, and funnel this absorbed energy to the PS-II reaction centers with efficiencies greater than 95%.

Phycobilisomes were discovered more than 30 years ago by Elizabeth Gantt and Sam Conti 1,2 in the outer thylakoid layer of the red alga Porphyridium cruentum. Phycobilisomes are present in large quantities in cyanobacteria and red algae, and may amount to as much as 50% of the soluble protein of the cell. Intact phycobilisomes can be readily dislodged intact from the membrane with a detergent such as Triton

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X-100 in a highly-concentrated phosphate medium to preserve their structural integrity, and then isolated by differential centrifugation on a sucrose-density gradient. The morphology of phycobilisomes in photosynthetic organisms can be recognized in, e.g., an electron micrograph of the thylakoid membrane, as shown in Fig. 2 (A) for the red alga Rhodella violacea, as shown in Fig. 2 (A). The view is of a cross-sectional cut perpendicular to the thylakoid membrane, where the phycobilisome granules are seen as semicircular aggregates. In some areas of the micrograph, the small rods radiating from the phycobilisome core can be seen (see inside of small circle). The electron micrograph at greater magnification in Fig. 2 (B) shows in more detail the structure of isolated phycobilisomes from the red alga Porphyridium cruentum, with each semicircular structure, ~40 nm wide, seen as consisting of a triangular core with six rods radiating from it. Fig. 2 (C) shows a model representing the thylakoid membrane of a cyanobacterium or a red alga, consisting of photosystems I and II interconnected by the cytochrome complex, and the ATP synthase, The phycobilisomes are seen as attached to the stromal surface at the PS-II reaction-center core complex.

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Phycobiliproteins and Phycobilisomes

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I. Phycobiliproteins Phycobiliproteins, such as phycocyanin (PC), phycoerythrin (PE), et al., are the building blocks of phycobilisomes in cyanobacteria and red algae, respectively. The chromophores in all cases have been found to be open-chain tetrapyrroles known as “phycobilins.” The structure of the four major types of phycobilins, namely, phycocyanobilin (PCB), phycoerythrobilin (PEB), phycourobilin (PUB), and phyobiliviolin (PBV), are shown in Fig. 3. The manner in which these various phycobilin pigments are covalently linked to their respective polypeptides is also depicted in the figure. The relationship between the spectral properties and structure of phycobiliproteins will be discussed first. We will then examine the supramolecular assemblies of phycobilisomes, how they are assembled from the phycobiliproteins, and how electronic excitation energy is transferred between them. Since the supramolecular assemblies of phycobilisomes are major light-harvesting complexes in both cyanobacteria and red algal cells, research on phycobiliproteins and phycobilisomes has been very active and extensive, and the deciphering of the experimental results to obtain an understanding of the detailed structure of phycobilisomes has been an exciting endeavor in the study of biological structures.

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I.A. The Phycobilin Pigments Unlike the chlorophyll-protein complex, where the pigments can usually be extracted with organic solvents, the extraction of the chromophores in phycobiliproteins require rather harsh chemical treatments as they are covalently linked to the proteins. Usual treatments include the use of 12 N HCl (or HBr) in or refluxing in methanol. From each of the phycobiliproteins, one can isolate the red-absorbing, phycobilin PCB and the blue-absorbing phycobilin PEB. The phycobiliprotein, R-phycoerythrin (RPE) [R for Rhodophyta], contains not only PEB, but also the 498 nm-absorbing (red) PUB. The protein phycoerythrocyanin (PEC) contains a fourth chromophore phycobiliviolin (PBV), which has also been called PXB [X implying the structure was at the time unknown], absorbing at 550-600 nm. The A-ring of each of the four phycobilins is bound to a cysteinyl residue of its respective polypeptide via a thioether linkage [see Fig. 3]. Phycocyanobilin may additionally be bound through ring-D, in some species. Phycoerythrobilin and phycourobilin may form two covalent bonds through both the A- and Drings with two cysteinyl residues ten amino acids apart (Cys-51 and Cys-60) in the polypeptide chain. In C-PC, two PCBs are bound to one polypeptide subunit, one through the A-ring and the other through the D-ring (not shown). Some phycobilins are isomers in a sense, differing only in the arrangement of the double bonds. For example, in group is linked to C18, but in PEB it was a group before linking a cysteine residue; there is also a difference in conjugation at C15 and C16. Note that small differences in the chromophore structure can produce a wide range of spectral properties in the phycobiliproteins. Isolated phycobiliprotein chromophores can assume two conformations: a porphyrin-like, helical geometry which absorbs mostly in the shorter wavelength region (300-400 nm), or an extended conformation which absorbs predominantly in the longer wavelength region (600-700 nm). Thus, a long-wavelength absorption by a phycobiliprotein indicates the presence of a phycobilin chromophore in the extended conformation.

I.B. Structure and Spectral Properties of Phycobiliproteins The two major polypeptides making up each phycobiliprotein are the so-called and subunits, with molecular weights of 17 and 18 kDa, respectively, and each consisting of 160-180 amino acids. Some phycobiliproteins, the phycoerythrins, contain an additional, 30-kDa subunit.

As depicted in Fig. 4, the and subunits in vivo form a monomer, and three monomers then link head-to-tail to form a trimer, which in turn combine with another trimer to form a hexamer. The

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and subunits of C-PC, for example, contain one and two PCB chromophores, respectively [see Table 1]. The trimer has the form of a disk 3 nm thick and 12 nm in diameter, with a 3.5-nm diameter hole in the center. In an unpurified extract of cyanobacteria or red algae, one may observe under an electron microscope not only the trimer disks but 6-nm thick hexamer disks as well, the hexamer disks clearly showing a faint line demarcating the two 3-nm disks.

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Depending on the species, phycobiliproteins may contain different types and numbers of phycobilin molecules linked to the protein subunits. Table 1 lists phycobiliproteins arranged into four classes based on chemical and spectroscopic properties. They are allophycocyanin (APC) absorbing between 650 and 670 nm, phycocyanin (PC) absorbing between 620 and 655 nm, phycoerythrocyanin (PEC) absorbing in the vicinity of 570 nm, and phycoerythrin (PE) absorbing between 540 and 570 nm. The corresponding wavelengths of the fluorescence peaks of the various biliproteins are listed in the last column. The precise wavelength value of absorption or fluorescence varies to some extent with any given species in a class. The wide variety of ways in which the subunits and the phycobilins are found combined in phycobiliproteins can be illustrated by three examples. In APC, each and subunit contains one PCB chromophore for a total of six for the trimer. In C-PC, each subunit contains one PCB chromophore and each subunit contains two, so a C-PC trimer contains altogether 9 PCBs. In R-PE there are six and six subunits and one protein subunit, i.e., With each subunit in R-PE containing two PEBs, each subunit two PEBs and one PUB, and the one subunit containing two PEBs and two PUBs, there is a total of 34 phycobilin molecules per Phycobiliproteins are water-soluble, slightly acidic with a pI ~ 5, and stable over a wide range of pH, from 5 to 9. All biliproteins are functionally and phylogenetically related, the different phycobiliproteins being thought to have descended from a common ancestor gene by gene duplication and subsequent mutations. The absorption spectrum of the monomer is essentially equal to the sum of absorptions of its components. On the other hand, the fluorescence quantum yield of the monomer is much higher than that of any of the separate components. These results suggest that association of the and subunits to form an monomer results in a reduction of the flexibility of the molecular skeletons of the bilins without altering their environment in any other spectroscopically significant way, as it is well known that a less flexible chromophore conformation inhibits the excited state from following alternative de-excitation pathways such as internal conversion. In the case of a denatured phycobiliprotein, the absorption is sharply decreased in the visible and it is non-fluorescent. However, noticeable changes in the absorption spectra occur upon formation of higher aggregates. For example, in C-phycoerythrin the wavelengths of maximum absorption in L and L (L standing for “linker” polypeptide; see Section II.A. below) are progressively red-shifted, with the absorption maxima appearing at 575, 577, 578 and 581 nm, respectively. With the formation of higher aggregates, not only the absorption spectrum, but such properties as the fluorescence emission spectrum, fluorescence lifetime and quantum yield, and circular dichroism, are also affected. The absorption and fluorescence spectra of the four representative phycobiliproteins are shown in Fig. 5: allophycocyanin from the filamentous cyanobacterium Anabaena variabilis, R-phycocyanin and Bphycoerythrin from the unicellular red alga Porphyridium cruentum, and R-phycoerythrin from the red alga Gastroclonium coulteri. The absorption spectra of phycobiliproteins cover a large part of the visible region, from ~490 nm to 670 nm. This wide range of color is actually generated by relatively small variations in the type and number of chromophores linked to the polypeptides, while their intense color can be accounted for by the large number of phycobilin chromophores they contain. For instance, phycoerythrin contains 34 chromophores in a disk 12 nm in diameter and 3 nm thick, giving an effective chromophore density of ~0.2 M. Each chromophore has a maximum molar extinction coefficient of at its absorption maximum in the visible region Phycobiliproteins are highly fluorescent, with a quantum yield between 0.3 and 0.9, which does not vary markedly with temperature. The emission intensity is constant over a broad range of pH and it is not known to be quenched by many biomolecules. These properties have in fact made phycobiliproteins very

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useful as fluorescent probes3: Conjugates formed from phycobiliproteins attached to certain small molecules with prescribed binding specificity are known as “phycofluors,” and are used as orange- or redemitting fluorescent probes in immunoassays and fluorescence-activated cell sorting.

I. C. Crystal Structure of Phycobiliprotein Subunits X-ray crystallographic analysis of C-phycocyanin from Mastigocladus laminosus (a thermophilic cyanobacterium) shows the phycobiliprotein to be a doughnut-shaped trimer [see Fig. 7 below]. The and subunits have tertiary structures that are similar to one another, consistent with their sequence

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homology. Fig. 6 (A) shows a stereo view of the structure of the subunit. The subunit, like the subunit, consists of a chain of eight interconnected helices, which may be represented as N(X,Y,A,B,E,F,G,H)-C. Helices A,B,E and F are nearly coplanar and tilted ~50° relative to helices G and H. Helices A, B, E, F, G and H together form the globular part of the subunit. It is worth noting that the arrangement of the helices in this globular portion of the and subunits is similar to that of the globins (hemoglobin and myoglobin). In fact, the lettering of the subunit helices has been adopted from that used for the globins, which have two additional helices, labeled C and D. these divergent evolutionary species,

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one a light-harvesting and one an oxygen-binding protein, are obviously phylogenetically related, and most likely derived from one common ancestor gene.

Each subunit contains two phycocyanobilins (PCB), one covalently bound to a cysteinyl residue at position 84 in helix E and the other to a cysteinyl residue at position 155 located on the loop between helices G and H [see stereo view in Fig. 6 (A)], while each subunit contains only one PCB covalently bound to the cysteinyl residue at position 84. Accordingly, these chromophores are designated as 155 and respectively.

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As noted in Section I.A, each of the chromophores bound to the polypeptide is in its extended conformation, that is stabilized by H-bonding to amino acid residues, with slight structural differences accounting for the different spectral properties of the chromoproteins. Although the degree of homology between subunit polypeptides from various cyanobacteria and red algae ranges from 21 to 67%, the cysteine residues at positions and (as well and used for linking phycobilins through their A- and D-rings are highly conserved. In each subunit, the helices X and Y at the N-terminus ofthe subunit form a U-shaped protrusion [see Fig. 6 (A)]. In the monomer, this protrusion is very close to helices a and e ofthe subunit [see Fig. 6 (B)]. The interaction between these protrusions determines the configuration of the monomer, whose overall shape resembles that of a “boomerang.” In the representation of monomer in Fig. 6 (B), uppercase letters are used for the helices ofthe subunit, with the lowercase letters for the subunit. Three monomers bind head-to-tail to form a doughnut-shaped trimer, [see Fig. 7 (A), left], 12 nm in diameter and 3 nm high, with a center-hole 3.5 nm in diameter. The interaction between two monomers leading to trimer formation is unlike the interaction between the and subunits inthe monomer itself. In the trimer, the loop between the B and E helices in the subunit of one monomer fits into a pocket formed by the X and Y helices of the subunit and the e and f helices of the subunit of the next monomer. The trimer has thus been likened to a “water wheel,” with the subunits forming the tips of the “paddles” [see Fig. 7 (A)]. The phycobilin pigments in the trimer are arranged in a circle; the chromophores are in the paddles ofthe wheel, the chromophores are near the periphery, and the chromophores closer in toward the center hole of the trimer [see Fig. 7 (A), right]. In the monomer, the center-to-center distance between the and chromophores is ~36 Å. In the trimer, the corresponding distance between the chromophore of one monomer and the chromophore ofthe adjacent monomer is about 22 Å. Interaction between the latter two chromophores at this close range accounts for the greater absorbance in the visible in the trimers compared to that in the monomers. Furthermore, since is known to be a sensitizing pigment, i.e., an energy donor and a fluorescing pigment, i.e., an energy acceptor, an approach this close between a sensitizing chromophore in an subunit and an fluorescing chromophore in a subunit would be expected to result in efficient energy transfer, as is observed. The structure ofthe hexamer formed from two rotationally staggered trimers is shown in Fig. 7 (B). As seen in the stereo view in Fig. 7 (B), left, the subunits in one trimer are on top of the subunits of another. Formation of a hexamer from two trimers face-to-face would involve mediation by the subunits. In the stereo view in Fig. 7 (B), left, the subunits are drawn in heavier lines and the subunits in thinner lines. A sketch of the six monomers (twelve subunits) is shown in Fig. 7 (B), right, where the numbers denote the monomers and the Greek characters and subunits.

II. Phycobilisomes As seen earlier in Fig. 2, a phycobilisome is a fan-shaped set of supramolecular assemblies, consisting of a triangular core with six short rods radiating outward, with each phycobilisome being attached to the thylakoid membrane through the core complex. In crude extracts of cyanobacteria, these short rods are seen in electron micrographs to have a diameter of ~12 nm, as expected for stacks of hexamers, with clearly marked divisions 6 nm apart along the rod and faint divisions midway between. In a medium of high ionic strength, in vitro hexamers can build up into short rods. However, such aggregation appears to require the presence of a so-called “linker” polypeptide (symbol “L”), which apparently serves as a kind of “glue.” Since it has been shown that phycobiliproteins purified to strict homogeneity cannot form these phycobilisome subassemblies. The presence ofthese linker polypeptides in

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cyanobacteria was first established by Nicole Tandeau de Marsac and Germaine Cohen-Bazire4 in 1977. Up to that time, it had been generally assumed that the phycobiliproteins alone could account for all the proteins in phycobilisomes. However, after examining phycobilisomes isolated from eight different cyanobacteria, these authors found that a small number of mostly colorless polypeptides, with molecular weights higher than any of the biliprotein subunits, amounted to ~15% of the total protein. Among these generally colorless proteins, those of the highest molecular weight (70-120 kDa) were found in the washed membrane fraction of the cell, which suggested at first that they are associated with the thylakoids and probably serve as a link between the phycobilisomes and the membrane.

II.A. The Linker Polypeptides As noted above, most linker polypeptides are colorless proteins, but some also contain phycobilin chromophores, endowing them with the ability to harvest light as well as apparently aid in the assembly of the phycobilisomes. While phycobiliproteins are acidic and hydrophilic, most linker polypeptides are basic and all are highly hydrophobic, electrostatic forces may also be an important factor in their function. With the recognition of the linker polypeptides, phycobilisome polypeptides may be classified into four groups according to their molecular weights as shown here, with the linker symbol, L, subscripted to indicate the specific linker function: 1. 2. 3. 4.

Group (1) includes the linker polypeptides, involved in the attachment of phycobilisomes to the thylakoid membrane, with the subscript “CM,” referring to its role as a linker at the core-to-thylakoid membrane junction. Each of these high-molecular weight polypeptides, which are found remaining in the thylakoid fraction after the phycobilisomes were removed by washing, contains one phycocyanobilin chromophore. The linker polypeptides and in Group (2) are either involved in assembling the biliproteins to each other in the rod, or in attaching the rod to the core. Group (3) includes the and subunits of phycobiliprotein as well as an 18.3-kDa polypeptide and possibly the subunit of APC. The 18.3-kDa polypeptide contains one phycocyanobilin and is the core subunit present in the phycobilisomes of most species. Since its sequence is homologous to that of the subunit of AP-B (allophycocyanin-B), the 18.3kDa polypeptide has often been abbreviated as Group (4) includes linker polypeptides present in the core complex, and are denoted by the symbol From various in vitro experiments, a picture has evolved to suggest a possible mechanism of how the linker polypeptide reacts with the biliprotein subunits to assemble the substructures of the phycobilisomes. Components and subassemblies of various phycobiliproteins together with their generic designations are shown in Fig. 8 (A). The various linker polypeptides are distinguished from each other by superscripts referring to their molecular weights. In an early step in the assembly of a rod, as represented by reaction (1) in Fig. 8 (B), the linker polypeptide acts as a primer to induce the formation of the trimer-linker complex, Under suitable conditions, the trimer-linker complex can serve as a template primer for the formation of a hexamer-linker complex, as shown in reaction (2) where two trimers are associated face-to-face [see layering in Fig. 7 (B) right]. Note that a large portion of the linker polypeptide is buried within the trimer formed in reaction (1), leaving only a small segment of the linker projecting from the face of the trimer. This small projection has been assumed to be the domain that links one hexamer with another linker complex, as shown in reaction (3) where this projecting segment links a with a to form a rod subassembly.

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II.B. Spatial Structure of Phycobilisomes in Anabaena variabilis When cyanobacterial or red algal cells are broken by osmotic shock in a low-salt solution, the biliproteins and phycobilisome subassemblies are released and solubilized. The size of the subassemblies depends on such conditions as pH, ionic strength, temperature, protein concentration, presence or absence of detergents. However, phycobilisomes can be isolated intact by breaking the cells in a moderately concentrated (0.65-1.0 M) Na/K phosphate buffer (pH 7-8) containing 1% or less Triton-X l00. After a short period of incubation intact phycobilisomes are released from the broken cells, aided by detergent action. The phycobilisomes may then be separated and purified free of chlorophyll by centrifugation in a sucrose density gradient of the same medium. The phycobilisome core is the site of attachment to the thylakoid membrane. Hemidiscoidal phycobilisomes, shown earlier in Fig. 2, are present in both cyanobacteria and some red algae, while hemiellipsoidal phycobilisomes are found only in certain other red algae. The phycobilisome in the thylakoid-less cyanobacterium Gloeobacter violaceus is present as a bundle of six rods; in the phycobilisome of Synechococcus 6301, the core (attached to the thylakoid) has only two short rods attached. The phycobilisome of Anabaena variabilis, shown schematically in Fig. 9, consists of six rods, each rod made up of 2-4 hexamer disks and attached to a core unit consisting of four trimer disks (fourth disk not represented in Fig. 9). Partial dissociation of some of the hexamer disks from the rods and subsequent examination by immuno-electron microscopy of the fractured phycobilisome structures that still remain bound to the membrane confirmed the general expectation that the shortest-wavelength absorbing PECs are present in the peripheral disks, while the longer-wavelength absorbing phycocyanin is present in disks proximal to the core, and the still longer-wavelength absorbing APCs and AP-Bs and a 120-kDa linker polypeptide, are present in the core. This order is consistent with the required electronic energy transfer properties for these disks, i.e., the wavelength of peak absorption shifts toward the red as one moves from the peripheral disks toward the core, and energy transfer is toward the core following the pathway,

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Phycobilisome rods of Anabaena variabilis contain mostly phycocyanin (PC), a minor amount of phycoerythrin (PE), and four linker polypeptides with molecular weights of 27, 29, 30.5 and 32.5 kDa. The phycocyanin and phycoerythrin content depends on culture conditions, such as light quality, temperature, etc. The order ofphycobiliprotein complexes along the rod is indicated for Anabaena variabilis in Fig. 9; the experimental confirmation of this sequence will be described below. Extensive fractionation of the dissociation products from A. variabilis phycobilisomes led to the isolation of three PC complexes, and and one phycoerythrocyanin complex With such trimer-linker complexes available, in vitro reconstitution experiments could be performed to help clarify the sequence of the components in the rods and how the linker polypeptides mediate rod assembly. As already shown in Fig. 8, a large portion ofthe linker polypeptide is buried inside the trimer, and only the small segment ofthe linker that is projecting outside the trimer is apparently able to link with the next trimer or hexamer. Limited tryptic degradation ofthe linker polypeptides and still associated with their respective biliprotein complexes, resulted in and respectively, i.e., 4.5- and a 6-kDa portions, presumably belonging to the exposed domains, were removed by tryptic degradation. These results are shown in Fig. 10 (A). As shown in Fig. 10 (B, 1), after the 4.5-kDa portion ofthe linker polypeptide had been removed from the original trimer complex, leaving its non-linker side (left face in the figure) is apparently unaffected as it can still attach another trimer to form a hexamer. However, without the projected domain this hexamer cannot attach to its right, as illustrated earlier in Fig. 8 (B, 3). It was also shown that with the projection still intact, apparently cannot attach to its right either. These results suggest that demands a specific and restricted location in the assembly, namely, the position nearest the core.

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When the exposed portion of is removed in a parallel experiment to form the hexamer by tryptic degradation, its left face remains unaffected, as the left face of the resulting is apparently able to continue to attach hexamers as shown in Fig. 10 (B, 2). Results from this experiment are consistent with the conclusion arrived at earlier, namely that or its truncated product, has to be at the terminal position, since its projection probably serves the function of attaching the rod to the core complex. A similar finding also confirmed to be at this location in a rod subassembly in Synechoccocus 6301. Lundell, Williams and Glazer5 reported earlier that in a solution containing the 27- and 33-kDa linker polypeptides and phycocyanin, the length of rods formed is inversely proportional to the ratio of the 27-kDa to the 33-kDa polypeptides, indicating that most likely occupies the terminal position where it inhibits any further lengthening of the rod.

II. C. Spectral Properties of Phycobilisomes As the supramolecular structure of phycobilisomes is formed, the absorption in the visible region is further enhanced. Apparently, the absorption is influenced by interaction of the chromophore with the protein environment, resulting also in a significant red shift of the absorption bands and a higher fluores-

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cence quantum yield. An additional influence is exerted by the linker polypeptide, as it functions not only as a linker but can also modulate the spectral properties of the biliprotein to which it is attached. It is obvious that for radiant energy absorbed by any one of the chromophores to be transferred efficiently along the rod toward the core and eventually to the chlorophyll in the thylakoid complex, it is necessary that the hexamer-linker complexes along the rod have appropriate spectral properties. Since all three hexamer disks in some rods may contain only PC, its spectral properties must be appropriately modulated to fulfill the condition for directional energy transfer. In Synechococcus 6301, for example, the spectral properties of the three PC hexamer disks along the rod are indeed modified by their linker polypeptides from the outer end of the rod to the core, as shown here from left to right:

This trend in spectral properties promotes energy transfer in the proper direction. Therefore, linker polypeptides not only have the function of linking biliprotein complexes together and controlling the position of the hexamer units along the rod, but also of modifying the spectral properties of the linked hexamers in order to achieve efficient energy transfer toward the core. In general, a phycobilisome contains several kinds of biliproteins, and its absorption spectrum reflects the contribution of each. On the other hand, as a result of energy transfer among the tightly coupled biliproteins in the phycobilisome, fluorescence is expected to originate mostly from the longest-wavelength component. As illustrated in Fig. 11 (A), phycobilisomes from cells of different species can have quite different absorption spectra. For instance, the absorption spectrum of a Porphyridium cruentum phycobilisome shows absorption bands characteristic of the component chromoproteins PE, PC and APC at 545-565,620 and 650 nm, respectively, while Anacystis nidulans only shows the absorption band of PC and a shoulder from APC. But, as shown in Fig. 11 (B), in spite of the quite different biliprotein

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composition, the fluorescence-emission spectra of both species appear nearly the same, each consisting of just a single peak at ~675 nm. This shows that energy transfer in both phycobilisoines is very efficient. Generally, therefore, with all phycobilisomes, irrespective of algal source or protein composition, as long as the structure provides tight energy coupling, the fluorescence emission spectrum appears almost always to originate from the terminal chromoprotein, adjacent to the core. The presence of fluorescence emission bands of shorter wavelengths would surely mean an inadequate energy coupling in the phycobilisomes, thus reducing the efficiency ofenergy transfer to the terminal point. On the other hand, when phycobilisomes are bound to the thylakoid membrane, the fluorescence of phycobiliproteins is expected to be low, because all the energy from phycobilisomes is channeled to chlorophylls in the thylakoids.

II.D. Energy Transfer among Chromophores in Anabaena Variabilis Phycobilisomes Brody and Rabinowitch6 and Tomita and Rabinowitch7 pioneered, more than forty years ago, excitationenergy transfer measurements in the subnanosecond range, from phycobilins to chlorophyll a in intact cells of several algae, using fluorescence lifetime studies. Energy transfer in phycobilisomes on the picosecond time scale was first examined, also more than twenty years ago, by Porter, Treadwell, Searle and Barber8 in intact cells of the red alga Porphyridium cruentum. They used a 565-nm pulse to selectively excite B-PE and found the fluorescence risetimes for R-PC (at 636 nm), APC (at 660 nm) and chlorophyll (at 680 nm) to be 12,24 and 50ps, respectively. These results were interpreted as evidence for a sequential energy transfer, i. e., Further insights into the mechanism of energy transfer in phycobilisomes were obtained by using picosecond measurements with even better time resolution as well as improved spectral resolution through the use of samples maintained at low temperature. We present here the study of Mamoru Mimuro and coworkers9 on time-resolved fluorescence spectra of both phycobilisomes and the APC-core complex isolated from Anabaena variabilis (M-3) cells. The phycobilisome rods in the M-3 cells contain PC only, thus simplifying the analysis. The APC core complex, isolated by partial dissociation of the phycobilisome, was useful for examining energy transfer restricted to the core portion only. Measurements were made on samples at 77 K using a 6-ps, 580-mn pulse from a mode-locked argon laser for preferential excitation of C-PC. Fig. 12 (A) shows fluorescence spectra of phycobilisomes recorded at different times after excitation. Initially a small emission band at 620 nm and a major band at 640 nm appeared, the latter shifting to 645 nm with time. The 645-nm emission due to PC reaches the maximum intensity at ~40ps, while the fluorescence peak near 660 nm from APC begins to appear at ~50ps. This 660-nm band shifts to 665 nm with time and reaches a maximum intensity at ~200ps, and then decays rapidly after that. Concomitant with the decay of the 665-nm emission is a rise of a 685-nm emission from the terminal emitter. At ~1 ns (see the 932-ps spectrum), the 645- and 665-nm bands have almost entirely decayed, and only the 685-nm band remains. These results are consistent with a sequential transfer of excitation energy from the rod to the core, i.e., terminal emitter. A more detailed picture of the energy-transfer sequence was revealed by additional kinetic analysis. Mimuro et al.9 obtained rise and decay curves for the individual chromophores by deconvolution of the time-resolved fluorescence spectra based on the relative intensities obtained by photon counting. The timeresolved fluorescence spectra could be computer-fitted to the reported emission spectra of nine major chromophores; four belonging to the rod PC and the remaining five to the APC-core complex. Deconvolution of the initial spectrum, when PC fluorescence is at a maximum, resolved four PC chromophore components with emission bands at 623,639,644 and 652 nm [see Fig. 12 (B)]. In the 665 and 685 nm regions the red shift of the maxima with time could be accounted for by two component bands

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each, namely the 660- and 666-nm bands belonging to APC and the 680- and 686-nm bands belonging to the subunit ofAP-B and the anchoring linker polypeptide, respectively. However, a best fit of the curves requires an additional band at 673 nm, which was attributed to the 16.2-kDa linker polypeptide in a core disk. The emission bands of the core components were further confirmed by deconvolution of timeresolved spectra obtained from the isolated APC-core complex, where interference by PC emission would be absent. Results obtained with the isolated APC-core complex agree well with those obtained with the phycobilisome, except an extra component at 673 nm is required for best curve fitting. The chromophore responsible for each fluorescence kinetic curve is assigned the letter “F” followed by the peak wavelength of the emission peak. Identification of the fluorescent chromophores, based on the previous measurements of Mimuro, Füglistaller, Rümbeli and Zuber10, showed that and and chromophores of PC rods emit at 622, 638 and 642 nm, respectively, and that energy transfer proceeds from to and then from to The fluorescent species (F) and the responsible chromophore belonging to each kinetic curve are listed on the right margin of Fig. 12 (B). For each kinetic curve the maximum intensity is indicated by a tick mark. The times at which the maximum intensity of each of the four fluorescence components originating from PC are achieved are consistent with a sequential energy transfer along the long axis of the phycobilisome rods toward the core. Energy flow among the individual chromophores of a phycobilisome rod is illustrated in Fig. 12 (C). From the chromophores identified, the fastest decaying is the one observed at 623 nm, its rapid decay most likely due to energy transfer from to or to another in an adjacent trimer [see Fig. 12(C), top]. Energy transfer between in one monomer and in an adjacent monomer is known to occur in less than 1 ps, and thus not detectable with the time resolution available. The F639 component has been interpreted as a reflection of a shift in the excitation equilibrium between the and chromophores, i.e., a change in the difference between the rate constants for forward and reverse energy transfer. The decay curves for F644 and F652 are nearly the same, and energy transfer between them is unlikely. It was concluded that the main energy flow in the rod occurs via the and chromophores in dual pathways along the rod axis, the chromophores functioning as an energy pool in rapid equilibrium with the chromophores. F652 most likely originates from (* indicating the presence of a linker to the polypeptide subunit) located in the trimer that serves as the energy donor to the adjacent core complex. The assignment of the fluorescence species in the core complex appears more complicated. Five fluorescence components, F660, F666, F673, F680, and F686 have been resolved [see Fig. 12 (B)]. The maximum intensities of the five kinetic curves do not show a straightforward sequential tendency. For instance, energy transfer from F666 to F673 does not appear to occur [see Fig. 12 (B)]. F660 and F666 have been assigned to the APC: without and with a 10-kDa linker polypeptide, respectively. It was previously reported by Mimuro, Lipschultz and Gantt11 that in Nostoc phycobilisomes an F680 attributed to an AP-B subunit and an F686 attributed to the 94-kDa anchor polypeptide are independent terminal emitters. Similarly, kinetic data in Fig. 12 (B) show that the rise of F686, rather than F680, corresponds to the decay of F673. These results led Mimuro et al.9 to propose two separate energy-transfer pathways in the core complex: one, corresponds first to a transfer from without a linker polypeptide to a 16.2-kDa linker polypeptide and then to the anchor polypeptide; the other, corresponds to a transfer from (with a linker polypeptide) to Mimuro et al.9 further pointed out that the core structure involving two independent, energy-transfer pathways as proposed above in Fig. 12 (C) is in basic agreement with the assembly model of GlazerR4, R5.

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For further reading: R1. WA Sidler (1995) Phycobilisome and phycobiliprotein structures. In: DA Bryant (ed) Molecular Biology of Cyanobacteria, pp 139-216. Kluwer R2. E Gantt (1986) Phycobilisomes. In: LA Staehelin and CJ Arntzen (eds) Encyclopedia of Plant Physiology, New Series, Vol. 19 pp 261-268. Springer Verlag R3. AN Glazer (1984) Phycobilisomes A molecular complex optimized for light energy transfer. Biochim Biophys Acta 768: 29-51 R4. AN Glazer (1985) Light harvesting by phycobilisomes, Annu Rev Biophys Chem 14, 47-77 (1985) R5. AN Glazer (1989) Light guides. Directional energy transfer in a photosynthetic antenna. J Biol Chem 264: 1-4 R6. H Zuber (1987) The structure of light-harvesting pigment-protein complexes. In: J Barber (ed) Topics in Photosynthesis, Vol 8. The Light Reactions, pp 197-259 R7. AR Holzwarth, Picosecond fluorescence spectroscopy and energy transfer in photosynthetic antenna pigments. In: J Barber (ed) Topics in Photosynthesis, vol. 8. The Light Reactions, pp 95-157. Elsevier R8. E M rschel and E Rhiel (1987) Phycobilisomes and thylakoids: The light-harvesting system of cyanobacteria and red algae. In: JR Harris and RW Horne (eds) Electron Microscopy of Proteins vol 6, pp 209-254. Acad Press R9. R Huber (1989) A structural basis of light energy and electron transfer in biology (Nobel Lecture), Angew Chem 28, 848-869 R10. M Mimuro (1990) Studies on excitation energy flow in the photosynthetic pigment system: structure and energy transfer mechanism. Bot Mag Tokyo 103, 233-253

References: E Gantt and SF Conti (1966) Ultrastructure of Porphyridium cruentum. J Cell Biol 26: 36-38 E Gantt and SF Conti (1966) Granules associated with the chloroplast lamellae of Porphyridium cruentum. J Cell Biol 29: 423-434 3. AN Glazer and L Stryer (1984) Phycofluor probes. Trends in Biochem Sci 9: 423-427 4. N Tandeau de Marsac and G Cohen-Bazire (1977) Molecular composition of cyanobacterial phycobilisomes. Proc Nat Acad Sci, USA 74: 1635-1639 5. DJ Lundell, RC Williams and AN Glazer (1981) Molecular architecture of a light-harvesting antenna. In vitro assembly of the rod substructures of Synechococcus 6301 phycobilisomes. J Biol Chem 256: 3580-3592 6. S Brody and E Rabinowitch (1957) Excitation lifetime of photosynthetic pigments. Science 125: 555 7. G Tomita and E Rabinowitch (1962) Excitation energy transfer between pigments in photosynthetic cells. Biophys J 2: 483-499 8. G Porter, CJ Treadwell, GFW Searle and J Barber (1978) Picosecond time-resolved energy transfer in Porphyridium cruentum. Part I. In the intact alga. Biochim Biophys Acta 501: 232-245 9. M Mimuro, I Yamazaki, N Tamai and T Katoh (1989) Excitation energy transfer in phycobilisomes at-196 C isolated from the cyanobacterium Anabaena variabilis (M-3): evidence for the plural transfer pathways to the terminal emitters. Biochim Biophys Acta 973, 153-162 10. M Mimuro, P F glistaller, R R mbeli and H Zuber (1986) Functional assignment of chromophores and energy transfer in C phycocyanin isolated from the thermophilic cyanobacterium Mastigocladus laminosus. Biochim Biophys Acta 848: 155-166 11. M Mimuro, CA Lipschultz and E Gantt (1986) Energy flow in the phycobilisome core of Nostoc sp. ( MAC ); two independent terminal pigments. Biochim Biophys Acta 933: 78-486 1. 2.


Chapter 15 The Primary Electron Donor of Photosystem II, P680, and Photoinhibition I.

The Primary Electron Donor of Photosystem II, P680 A. Absorbance Changes accompanying P680 Photooxidation B. The EPR Spectrum C. The Electrochemical Properties II. Photoinhibition in Photosystem II Consequence of the Strong Oxidizing Power of P680 + A. Two Pathways leading to Photoinhibition in PS-II Reaction Centers B. Elucidation of Photoinhibition in vitro with the D1/D2/Cyt b559 Complex C. Degradation of Protein D1 during Photoinhibition III. Possible Protective Function of Cytochrome b559 against Photoinhibition A. Some Properties of Cytochrome b559 B. Photooxidation of Cytochrome b559 C. Photoreduction of Cytochrome b559 D. Proposed Role for Cytochrome b559 in Protecting against Photoinhibition

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Photosystem II of the photosynthetic thylakoid membrane is activated only by light with wavelength measurably shorter than 700 nm. When the primary electron donor P680 is photoexcited it becomes oxidized and an “intermediary” electron acceptor a pheophytin molecule, is reduced: (The subject of will be discussed in detail in Chapter 17.) The couple has a very positive (i.e., oxidizing) redox potential enabling it to extract electrons (indirectly) from water, resulting in the splitting of water molecules into molecular oxygen, electrons and protons. The extra electron on is transferred further to the protein-bound plastoquinone molecule In photosystem II, the semiquinone of must first transfer an electron to the protein-bound plastoquinone before it can receive another electron from in a second round of photoexcitation. however, can accept two electrons in succession to form the fully reduced hydroquinone (see Chapter 16 for details). This reduced hydroquinone molecule is then exchanged with an oxidized plastoquinone molecule in the quinone pool in the membrane matrix, and this two-electron reaction cycle can then be repeated. The reduced plastoquinone eventually goes to reduce the photooxidized primary donor of photosystem I, resulting in the uptake and release of protons on opposite sides of the thylakoid membrane and thus providing an important mechanism for the storage of some of the absorbed solar energy. The sequence of electron-transfer events may be written as:

where the “double arrows” stand for other electron-transfer steps, which are the topics discussed elsewhere in the book.

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The electron used for restoring (i.e., reducing) ultimately comes from a water molecule, via a cluster of Mn atoms and a secondary electron-donor molecule called Stoichiometry requires four, one-electron transfer steps to evolve one molecule of oxygen from two molecules of water. The states generated by this four-step process are called the “S-states.” Oxygen evolution promoted by electrontransport through photosystem II may be written as:

In this chapter, we will look at how charge separation takes place in PS-II reaction centers after photoexcitation and at the properties of the PS-II primary electron donor P680. In the following chapter we will discuss the so-called “stable primary” electron acceptor and the secondary electron acceptor This will be followed by a discussion of the “intermediary” electron acceptor, the species that actually accepts the electrons from the photoexcited primary donor P680*. We adopt this sequence ofpresentation because the reduction of Q was experimentally more readily observed than that of and was quite naturally the first experimentally observed acceptor in the course of photosystem-II research. The importance of biological oxygen evolution can hardly be overemphasized from the standpoint of the biosphere, as practically all the earth’s supply of atmospheric oxygen comes from and is maintained by the regeneration processes carried out by algal and plant photosynthesis. Until recently, detailed knowledge of the mechanism of water splitting to release oxygen had been lacking but in the past few years some important experimental advances have provided valuable new insights. The subject of water oxidation will be discussed in Chapters 18 to 24, which include various other related topics, including the secondary electron donor to the photooxidized

I. The Primary Electron Donor of Photosystem II, P680 We may depict the PS-II reaction center core that has the minimum composition for light-induced charge separation simply as but we may extend the concept of the reaction-center core to include a secondary donor, and a secondary acceptor, Q, so that we can visualize the steps beyond the primary charge separation:

I. A. Absorbance Changes accompanying P680 Photooxidation The primary electron donor P680 of PS II was first characterized by Döring, Renger, Vater and Witt1 by means of flash-induced absorbance changes in spinach subchloroplasts enriched in PS II. The difference spectrum they obtained from 420 to 725 nm is shown in Fig. 1 (A). The rate of P680 bleaching was reported to be The portion of the signal with a decay time of ~200 was used for plotting the absorbance-change spectrum, which displayed major absorbance decreases at 435 and 682 nm and a minor one at 640 nm. Note that the term “P680” was first coined by Rabinowitch and GovindjeeR7 in their Scientific American article in 1965 and widely adopted in current photosynthesis literature for the primary donor but it was called by Döring et al. 1 . It should also be mentioned that since the strong PS II fluorescence generated by the excitation beam in the red region can interfere with the measurement of P680 absorbance changes, the monitoring beam was modulated at a high frequency so that the absorption-change signal could be extracted from the fluorescence background signal2. Döring et al.1 also reported that the P680 change can only be elicited by light at wavelengths below 700 nm.

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PS-II Primary Electron Donor P680 and Photoinhibition

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It was subsequently found by van Gorkom, Pulles and Wessels3 that P680 photooxidation is also accompanied by an absorbance increase in the 820-nm region, as shown by the “light-minus-dark” spectrum in Fig. 1 (B), obtained with digitonin-fractionated PS-II subchloroplasts. It can be seen from this figure that although there are no sharp features in the near-infrared region, there is nevertheless clearly an absorbance increase, with a maximum near 820 nm. Thus P680 changes can be monitored in this farred region without interference from fluorescence. However, it should be pointed out that this far-red absorbance increase is not exclusive to P680 since, for example, P700, the primary electron donor of photosystem I, also produces an absorbance increase in this spectral region when it is photooxidized. Using near-infrared light for monitoring P680 photooxidation does have some other advantages, the most important being that the monitoring light does not cause photochemical bleaching of the chloroplast particles. Furthermore, photodiodes emitting coherent, near-infrared laser light are readily available as monitoring-light sources. As is now known, P680 undergoes photooxidation not just in a few nanoseconds but actually in a few picoseconds. We will defer the discussion of the rapid kinetics of P680 photooxidation to Chapter 17. As mentioned above, the difference spectrum in Fig. 1 (A) is that of changes with a decay time of ~200 which are probably due to the back-reaction of with a reduced secondary electron carrier on the reducing side of photosystem II. Subsequently it was found that with greater time resolution two small components of decay in ~35 ns and ~30 respectively. These multiple decay rates were interpreted as due to recombination of two different populations of with other electron carriers, such as etc., in the PS-II reaction center: Although under normal, active oxygen-evolving conditions the rapid nanosecond decay components of photooxidized are difficult to detect and measure without adequate time resolution, the components arising from the reaction with the secondary donors may be retarded by various chemical treatments such as Tris or low pH. After any of these treatments, the predominant route of decay of is the slower recombination with in 200 at room temperature. At sufficiently low temperatures, photoinduced charge separation in PS II still takes place, but cannot react with even in chloroplasts capable of oxygen evolution at room temperature. Instead, most of the recombines with in ~3 ms. At low temperatures, a small portion of can oxidize cytochrome b559 present in the reduced state. Cytochrome b559 is a chloroplast component of yet unknown role; its properties and possible function will be discussed below under Section III. A representative absorbance-change signal for P680 photooxidation in steady light is shown in the inset of Fig. 1 (B). The rapidly decaying portion, which presumably represents the recombination of and was used to construct the difference spectrum. Thus, it should be recognized that the difference spectrum is a sum ofchanges due to both P680 and i.e., Changes in the red region probably belong to P680 only, while those in the ultraviolet region that has been examined belong primarily to In later work, Gerken, Dekker, Schlodder and Witt4 have been able to obtain a net difference spectrum representing changes in the blue region due to P680] exclusively, as shown in Fig. 1 (C). These workers, using an oxygen-evolving PS-II particle prepared from a cyanobacterium Synechococcus, applied chemical intervention so that after a net difference spectrum representing the change was obtained, a net difference spectrum representing only the reaction was obtained under a different set of conditions (details for obtaining these two difference spectra will be discussed in the next chapter). By subtracting the difference spectrum for the reaction from that for the reaction, a net difference spectrum belonging only to the reaction was obtained.

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It is of interest to note that although the reports quoted in Fig. 1 span a period of more than 20 years, the three results are in fairly good agreement. This is also a good example for illustrating progress in photosynthesis research, in terms of the purity and activity of the experimental samples available, the precision of techniques for acquiring the optical difference spectrum, and the increase in knowledge for controlling the reaction route. The difference spectrum of the reaction shown in Fig. 3 (C) encompasses the fruits of all these advances.

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I.B. The EPR Spectrum Since is normally re-reduced very rapidly by the endogenous secondary electron donor the EPR signal of would ordinarily be rather difficult to observe. Furthermore, the highly oxidizing can also extract electrons from pigment molecules such as chlorophyll and carotenoid as well as other electron carriers, and many early EPR spectra reported as originating from often included a significant contribution from Its EPR spectrum is very similar to that of having the same gvalue of 2.00 and, as shown later, only a slightly larger halfwidth, making it difficult to distinguish one from the other. In practice then, an EPR signal due exclusively to can be observed only when these potential electron donors are inhibited or blocked, or by using time-resolved detection methods, or by a combination of both. In Fig. 2 we present the results using both of these methodologies. In uninhibited oxygen-evolving PS-II subchloroplasts, photoinduced is rapidly reduced by in and thus no EPR signal for formation is observable if the response time of the measuring instrument is, say, 50 or greater. Under these conditions, however, the EPR signal of the secondary donor generated by the oxidation of by can be observed as a so-called II very fast) as its lifetime is relatively long. A detailed discussion ofthis signal will be found in Chapter 22.

276

If is added to PS-II subchloroplasts to abolish oxygen evolution by blocking electron flow from the secondary donor to photoinduced extracts an electron less rapidly from an alternate source, namely, the photoreduced primary electron acceptor Fig. 2 (A) shows the transient kinetics of a flash-induced, EPR signal change in the g=2 region, where the initial, fastrising component reflects the photooxidation of P680, i.e., and the portion rapidly decaying in ~200 reflects the reduction of by reduced The small and much more slowly decaying component presumably represents a slow reduction of some of the by other secondary donors. The spectrum shown in Fig. 2 (B) is constructed from the amplitude of the rapidly decaying component measured over a range of magnetic field values. The spectrum, with a g-value of 2.002 and a linewidth of 7-8 G, is attributed to Similar results were also obtained with a PS-II particle prepared from the cyanobacterium Synechococcus after treatment with a high concentration (0.6 M) of acetate to inactivate the components closely involved in oxygen evolving activity. Treatment with concentrated acetate is known to retard the reduction of by the secondary donor by several orders of magnitude, thus making the effective electron donor to this inhibition can be completely reversed by the removal of the acetate. On the other hand, if an “ADRY” agent such as FCCP is also added to the acetate-treated PS-II particles, reduction then occurs by a somewhat slower but still effective electron flow from the endogenous secondary donor Discussion of the secondary electron donors to and and the so-called “ADRY” agent (the term comes from “acceleration of the deactivation reaction of the water-splitting enzyme Y”) will be presented in Chapter 22. Figure 2 (C), trace (a) shows an EPR transient of an acetate-FCCP treated PS-II particle with an instrument-limited risetime and a decay time of 170 corresponding, respectively, to the oxidation of P680 to and the reduction of the latter by Transient (b) has a risetime corresponding to the decay time of but it decays in milliseconds; it can thus be attributed to An EPR spectrum constructed from the rapidly-decaying transients [the solid-line spectrum (a) in Fig. 3 (D)] therefore represents It has a g-value of 2.0027 and a linewidth of 8 G. The dashed-line EPR spectrum (b) in Fig. 3 (D) is constructed from the amplitude of the slowly-decaying component. This EPR spectrum has a zero crossing at g=2.0044 characteristic of a semiquinone and has the line shape of an SII-type signal (see Chapter 22). We now briefly discuss the question of whether P680 is a monomer or dimer. As mentioned in the introductory chapter, photosystem II closely resembles the reaction center of purple bacteria in its biochemical makeup. This similarity, therefore, naturally leads to the notion that the PS-II primary donor P680 might also have a dimer configuration as in the bacterial case, i.e., it is present as a special chlorophyll pair. However, the monomer-or-dimer question with regard to P680 has remained uncertain for some time. In regard to its optical absorption spectrum, it is unlike P700, whose wavelength maximum is appreciably red-shifted from that of the bulk chlorophyll and which was long thought to be a dimer. The wavelength maximum of P680 absorbance change is not as red-shifted, which would suggest that it is more likely a chlorophyll monomer. On the other hand, the EPR linewidth of 7-8 G, established by both measurements shown in Fig. 2, would suggest a dimer configuration for P680, as this linewidth appears to be narrowed from that of the monomer by a factor of (refer to discussion on bacterial reaction-center dimer in Chapter 4). In comparing the redox potential estimated for Davis, Forman and Fajer5 found it not far from that of the couple in vitro (whose redox potential has been estimated as high as +0.9 V), and much higher than that of the couple. This consideration led the authors to suggest that EPR linewidth alone cannot be used as a conclusive indicator for the composition of P680 in vivo. Instead, the authors suggested that P680 may be a ligated monomeric chlorophyll whose EPR and redox

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properties are modified by some specific interaction with the amino-acid residue(s) of the protein environment. There are also other data in conflict with an assumed dimer configuration. The EPR signal obtained for the spin-polarized triplet state of P680 formed by recombination from the light-induced state (for details see Chapter 17) has zero-field-splitting parameters identical to those of the triplet state of monomeric chlorophyll in vitro. Also the triplet-minus-singlet optical spectrum of P680 measured by the absorption-detected magnetic resonance (ADMR) technique does not support a dimeric structure for P680. The current status of the P680 dimer-monomer question may best be summarized by quoting the assessment of van der Vos, van Leeuwen, Braun and Hoff6 based on currently available evidence, “we conclude that P680 is functionally monomeric. This does not necessarily mean that only a single monomeric Chi a absorbs at 680 nm. Two Chl a molecules may contribute and may actually be located close to the position suggested by the primary donor dimer in the bacterial reaction center, but they are not excitonically coupled, and only one of them appears to serve as the primary donor of PS II.”

I. C. The Electrochemical Properties A precise redox potential for has not yet been directly determined but it must have a very high redox potential in order to be able to oxidize Since the redox potential associated with the water molecule is +0.82 V, and since there are additional electron carriers present between P680 and water, the redox potential of the couple must be considerably more positive (more oxidizing) than +0.82 V. In 1977, Jursinic and Govindjee7 and in 1979, Klimov, Allakherdiev, Demeter and Krasnovsky8, independently estimated the redox potential of the couple. The minimum photon energy needed to electronically excite P680 is known to be in the range 1.8-1.82 eV. Klimov et al. separately determined the redox potential of the couple in vitro to be V8 (see details in Chapter 17). They further took into consideration that the energy barrier between P680* and as estimated from the activation energy of delayed fluorescence to be 0.04-0.08 eV9. From these values, the redox potential of the couple was estimated to be V. 7 Jursinic and Govindjee , based on data derived from temperature dependence of delayed light emission, calculated the redox potential of P680 as between 1.0 to 1.3 V. Either of the redox-potential values should provide with ample oxidizing power to carry out the reaction, whose redox potential lies almost 300 mV lower. In Chapter 22 we will describe another method for estimating the potential of the couple by indirectly titrating the secondary electron donor as monitored by its EPR signal, the value estimated for by this method is about +1.2 V, close to values estimated by Jursinic and Govindjee7 and by Klimov et al.8.

II. Photoinhibition in Photosystem II – Consequence of the Strong Oxidizing Power of Light energy is the benefactor and the ultimate driving force for photosynthesis. But ironically, excessive light, particularly in combination with certain other stressful conditions, may be detrimental to photosynthetic activity. This phenomenon is known as “photoinhibition” or “photodamage.” Photoinhibition results in the destruction of pigment molecules and in the degradation and cleavage of the D1 protein [see Fig. 3 (A)]. Damage caused by too much light is usually mended by replenishing these degraded components through renewed synthesis and reassembly of the PS-II reaction-center complex. Photoinhibition is observed when the rate of repair is less than the rate of photodamage. Clearly, photoinhibition (photodamage) is likely to occur in photosystem II due to the extremely high oxidizing power of

278

In normal photosynthesis, light-induced charge separation occurs in both photosystems I and II and the flow of electrons from water by PS II is eventually used by PS I to reduce (and generate a trans-membrane proton gradient used for the synthesis of ATP). Under certain circumstances, with a redox potential possibly as high as +1.2 V, may, as previously noted, extract an electron from a nearby species other than the normal secondary donor, possibly a pigment molecule. Under high light intensity, it is also possible for to undergo double reduction, and thus be unable to accept an electron from a newly formed The result is that the separated charges in recombine. This recombination occurs partly by way of the excited singlet of P680 and partly by way of the excited triplet state of The lifetime of is sufficiently long that it can with high probability react with ground state triplet oxygen to form singlet oxygen, which is known to be damaging to pigment and protein molecules. Carotenoids present in the bacterial reaction center or in the light-harvesting chlorophyll-protein complex usually have a protective function by quenching any chlorophyll triplet. However, carotenoids present in the PS-II reaction center apparently do not perform this function directly. Presumably this is a consequence of the fact that carotenoids have to be sufficiently distant from P680 in order to avoid photooxidative destruction themselves.

II. A. Two Pathways leading to Photoinhibition in PS-II Reaction Centers As envisioned by Jim Barber and Bertil Andersson10, photoinhibition may follow either of two pathways: one when the acceptor side is impaired, called “acceptor-side induced,” and the other when the donor side is impaired, or “donor-side induced.” Telfer and BarberR5 have made use of the PS-II reaction-center complex D1/D2/Cyt b559 to elucidate these two pathways. The D1/D2/Cyt b559 complex [Fig. 3 (A)] is composed of the D1 and D2 proteins and the heterodimeric, heme-containing Cyt b559. The complex binds four to six chlorophylls (of which two make up P680 and the remainder are monomeric chlorophylls), two pheophytins and two (Car). The plastoquinone molecules and and the nonheme iron atom as well as the manganese atoms are apparently lost during the isolation of the complex. The tyrosines that serve as the secondary electron donors and are also no longer active. The inactive or absent species are shaded in the figure.

II.B. Elucidation of Photoinhibition in vitro with the D1/D2/Cyt b559 Complex Because it is devoid of the active, endogenous secondary acceptors and donors, the D1/D2/Cyt b559 complex can serve as an extremely valuable experimental system for the elucidation of the mechanism of photoinhibition. Since the complex contains the active primary donor and acceptor, it can catalyze secondary electron transport if both an artificial secondary donor such as diphenyl carbazide (DPC) or a quinone acceptor such as 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone (DBMIB) and an artificial secondary acceptor such as silicomolybdate (SiMo) or ferricyanide (FeCy) are provided, as shown in Fig. 3 (B). The Dl/D2/Cyt b559 complex, as prepared and without any additions, is already impaired on the acceptor side. Strong illumination causes charge separation, followed by charge recombination, and formation. When oxygen is present, interacts with oxygen to form the very reactive singlet oxygen, which in turn will proceed to attack the various reaction-center components [see the acceptor-limited reaction pathway in Fig. 3 (C)]. A donor-limited photoinhibition pathway in the D1/D2/Cyt b559 complex can be created by the addition of the artificial electron acceptor silicomolybdate (SiMo), as shown in Fig. 3 (D). SiMo is an efficient, exogenous secondary electron acceptor of photosystem II. After light-induced charge separa-

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tion, SiMo drains electrons from reduced pheophytin, and consequently, becomes photoaccumulated. As previously noted, the highly oxidizing can be very detrimental to various PS-II components. Photoinhibition following one or the other pathway is illustrated by the bleaching of various pigment molecules by the reactive species as shown in Fig. 4. In the acceptor-limited photoinhibition pathway, the reaction-center complex appears to be resistant to photodamage when oxygen is absent. As seen in Fig. 4 (A), absorbance changes measured in the absence of oxygen show negligible bleaching in the 680 nm region, but clearly a slow reduction of Cyt b559, presumably by reduced When oxygen is present, interacts with oxygen to form as confirmed by the results of several independent experimental results, including: (1) the shortening of the lifetime of from 1 ms in the absence of oxygen to 33 in the presence of oxygen11, and (2) the detection of singlet oxygen directly by both steady-state and time-resolved measurements of its phosphorescence at 1270 nm11. The absorbance difference spectra produced by illumination under the acceptor-limited condition at a photon flux density of~1000 for half a minute shows a bleaching at 417, 435 and 680 nm, indicating that the chlorophyll of P680 is being attacked initially. Under prolonged illumination for five minutes, the bleaching increases, and the band maximum also shifts slightly to the blue, indicating that the monomeric accessory chlorophylls are being attacked. There is also some spectral indication (e.g., absorbance decrease at 545 nm) that pheophytin molecules may also be attacked. There is no indication of Cyt b559 being reduced, however. Furthermore, appears quite stable toward photodamage in the acceptor-limited pathway either in the absence or presence of oxygen.

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Illumination under the donor-limited condition, either in the absence or presence of oxygen, produces absorbance changes quite different from those under the acceptor-limited condition. There is, e.g., a prominent irreversible bleaching of and the accessory chlorophyll absorbing at 670 nm [see Fig. 4 (B)]. As a result of the oxidation of by the secondary acceptor SiMo, the lifetime of is greatly increased and thus intensifying its destructive effect. Of course, this donor-limited photoinhibition can be readily prevented if an artificial secondary electron donor such as DPC or is also added.

II. C. Degradation of Protein D1 during Photoinhibition Accompanying the above-mentioned bleaching ofvarious pigment molecules during photoinhibition is a loss of photosynthetic activity and degradation of the D1 protein. Greenberg, Gaba, Mattoo and Edelman12 first found in 1987 a degradation product in the form of a 23.5 kDa fragment during PS-II photoinhibition, and concluded that it was formed by a proteolytic cleavage in the loop connecting the D- and E-helices of the D1 protein [see scissors in Fig. 4 (C)]. The pattern of protein degradation depends on whether photoinhibition takes place via the acceptoror donor-limited pathway. Studies using the RC complex D1/D2/Cyt b559 as the experimental system have shown that under the acceptor-limited condition, a polypeptide fragment is formed having an apparent molecular weight of 23-kDa and containing the N-terminus, and that its yield increases steadily with time of illumination. The appearance of this 23-kDa polypeptide fragment is accompanied by the bleaching of the 680 nm-absorbing chlorophyll, the extent of degradation being oxygen dependent, as expected for an acceptor-limited pathway. Another photo-degraded polypeptide fragment has a molecular weight of 10-kDa and contains the C-terminus [see Fig. 4 (C) bottom]. These results agree with the initial finding of Greenberg et al.12 that cleavage occurs in the DE loop of D1. When photoinhibition follows the donor-limited pathway, a 24-kDa polypeptide fragment containing the C-terminus and a 9-kDa fragment containing the N-terminus are produced, indicating that under the donor-limited condition, the D1 protein is cleaved in the loop connecting the A- and B-helices [see scissors in Fig. 4 (D)]. Similar to the pigment bleaching by the donor side-limited, photoinhibition pathway, the cleavage of D1 into a 24-kDa C-terminal fragment and a 9-kDa N-terminal fragment is also independent of the presence of oxygen. It is worth noting that the photoinhibitory condition can also cause degradation of the D2 protein. However, given normal light intensity, the D1 protein is apparently degraded at a significantly higher rate than the D2 protein. At present, the precise enzymatic mechanism of protein D1 degradation during photoinhibition is unknown. However, it is of interest to note that the cleavage sites of the D1 protein are quite specific in the two photoinhibition pathways. As seen from Fig. 4 (C) and (D), protein cleavage in the acceptor- and donor-limited photoinhibition pathways occurs on the stromal and lumenal side, respectively, of the protein (indicated by the scissors locations in the figure), and that both cleavage sites are located in the hydrophilic loops of the protein. Conformational changes in the protein as a consequence of pigment degradation, among other factors, undoubtedly play an important role in photoinhibition. The applicability of these two pathways to in vivo photoinhibition is to a large extent still an open question. Telfer and BarberR5 subjected normal intact leaves (of peas and wheat) to photoinhibitory conditions and reported the appearance of a 10-kDa C-terminal protein fragment, indicative of an acceptor-limited pathway but results consistent with a donor-limited photoinhibition pathway have not yet been experimentally demonstrated in vivo.

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III. Possible Protective Function of Cytochrome b559 against Photoinhibition

III.A. Some Properties of Cytochrome b559 Cytochrome b559 is an integral membrane protein of chloroplast thylakoids with an absorption maximum at 559 nm (hence its name). It is one of three redox heme proteins present in higher plants and algae, the other two being cytochrome f and cytochrome The molar ratio of Cyt b559 to the PSII reaction center is unity, as revealed by analysis of many types of PS-II subchloroplasts, including highly active, oxygen-evolving particles, as well as by analysis of other thylakoid membranes. Furthermore, while chloroplast cytochromes and f are part of the Cyt whose function is well understood, the physiological function of Cyt b559 remains largely unclear. Much speculation has been made over the years regarding the function of Cyt b559; in 1978, Warren Butler13 even proposed that Cyt b559 may be involved in the oxygen evolving apparatus. Since the physiological function ofcytochrome b559 is as yet unknown, it is difficult to fit it into any suitable spot for its presentation. We place the discussion of cytochrome b559 at this point, however, because it is known to be a component closely associated with photosystem II and presumably close to the major PS-II components P680 and Another reason is of more recent interest: Cyt b559 has been proposed by several groups of workersR5, R6, R7 as possibly having a protective function against photoinhibition. In the following we will first describe some physical and chemical properties of Cyt b559, and then show examples of its photoinduced reduction and oxidation in chloroplasts and subchloroplast particles. Finally we will briefly discuss its possible protective role against photoinhibition. Cytochrome b559 is bound to the photosynthetic membrane and can be present in either of two forms with different redox potentials (refer to Fig. 5 below). In freshly prepared chloroplasts, it has a redox potential of~350 mV and is thus designated “Cyt b559 (HP)” [HP for high-potential]. Cyt b559 (HP) can be readily reduced by hydroquinone. When fresh chloroplasts are variously treated, such as by aging or heating, or with chaotropic agents, detergents (e.g., Triton), trypsin, and Tris, Cyt b559 is transformed into a low-potential form, with a redox-potential value of ~0 to +50 mV, and designated as “Cyt b559 (LP).” This low-potential Cyt b559 is readily oxidized either chemically or photochemically. Cyt b559 is a heterodimer consisting of a 9-kDa with 83 amino-acid residues and a 4kDa with 39 residues, the two polypeptides joined by a heme through a pair of histidines, one on each polypeptide chain close to the N-terminus, as shown in Fig. 5 (A). Each polypeptide subunit contains one transmembrane helix, the N-terminal ends of both polypeptide subunits being located on the stromal side. Babcock, Widger, Cramer, Oertling and Metz14 investigated the structure of Cyt b559 obtained from both spinach and maize by characterizing its ligands and spin state by various spectroscopic methods, using the compound, bis(N-methylimidazole) iron protoheme, as a model. In both the oxidized and reduced proteins the heme iron is in the low-spin state. The authors concluded that the heme iron is ligated in its fifth and sixth (out-of-plane) coordination positions by histidines of the and tide subunits, as shown schematically in Fig. 5 (A). The EPRg-values of Cyt b559 (LP) in purified and membrane-bound forms are essentially identical to those of the model compound, indicating that the histidine planes are oriented parallel to each other, and that the heme normal is parallel to the membrane plane [see Fig. 5 (B), bottom]. However, Cyt b559 (HP) exhibits significantly different g-values from the model compound, which led Babcock et al.14 to suggest that the different thermodynamic properties of the LP and HP forms (i.e., the different redox potentials) and the different g-values may have their origin in differences in the stereochemistry of the histidine planes. As shown schematically in Fig. 5 (B), the histidine planes in their model of the LP- and HP-forms have different relative orientations: a parallel orientation in the LP-

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form and a nonparallel orientation in the HP-form. In other words, the heme ligand stereochemistry alters the g-values and influences the redox potential. This hypothesis may also explain the ability of the HP- and LP-forms to interconvert as a result of environmental (membrane) state changes. While various treatments such as aging, chaotropic agents, etc. convert the HP- into the LP-form, as mentioned above, it has been reported that the LP-form can also be converted into the HP-form by addition of lipid to the membrane.

III. B. Photooxidation of Cytochrome b559 Cytochrome b559 has no known physiological role, although it can undergo photoinduced oxidation or reduction depending on sample conditions. Irreversible, light-induced oxidation of Cyt b559 in lettuce chloroplast near liquid-nitrogen temperature was reported by Knaff and Arnon15 more than 30 years ago. As seen in Fig. 6 (A) top, Cyt b559 photooxidation is elicited by short-wavelength red light (664 nm) but not by far-red light (714 nm), clearly showing that the photooxidation is promoted by a PS-II reaction. If FeCy is first added, however, no absorbance changes which can be attributed to Cyt b559 occurs in the light, confirming that photooxidation had occurred in the absence of FeCy [see Fig. 6 (A), bottom]. It was subsequently found that only the low-potential Cyt b559 (LP) present in the reduced state can undergo low-temperature photooxidation.

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More recently Thompson and Brudvig16 examined a similar reaction in spinach chloroplast by EPR spectroscopy. In the active oxygen-evolution pathway, extracts electrons from the secondary donor more rapidly than from Cyt b559 or a chlorophyll molecule designated as It is thought that which is present in the reaction center, serves primarily as a link for energy transfer from the antenna to P680. These three species can be considered as competing electron donors to and only when donation by is significantly diminished in some way, will Cyt b559 and reduce to a significant extent [see Fig. 6 (A) and Fig. 7 (A)]. This is consistent with the low-temperature, irreversible photooxidation of Cyt b559 in lettuce chloroplasts as described above. These workers measured, over a range of temperatures, the yield of photooxidation of the alternate electron donors relative to the yield of photooxidation of manganese as shown in Figs. 7 (B) where the percentage of the maximum yield for each signal is plotted as a function of the illumination temperature. At low temperatures (

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